D12009.PDF

WORLD ORGANISATION FOR ANIMAL HEALTH
MANUAL OF DIAGNOSTIC TESTS AND VACCINES
FOR TERRESTRIAL ANIMALS
(mammals, birds and bees)
Seventh Edition
Volume 2
2012
This Terrestrial Manual has been edited by the
OIE Biological Standards Commission and adopted by
the World Assembly of Delegates of the OIE
Reference to commercial kits does not mean their endorsement by the OIE. All commercial kits should
be validated; tests on the OIE register have already met this condition (the register can be consulted
at: www.oie.int).
OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals
Seventh Edition, 2012
Manual of Recommended Diagnostic Techniques and Requirements for Biological Products:
Volume I, 1989; Volume II, 1990; Volume III, 1991.
Manual of Standards for Diagnostic Tests and Vaccines:
Second Edition, 1992
Third Edition, 1996
Fourth Edition, 2000
Fifth Edition, 2004
Sixth Edition, 2008
Seventh Edition: ISBN 978-92-9044-878-5
Volume 2: ISBN 978-92-9044-880-8
©
Copyright
World Organisation for Animal Health (OIE), 2012
12, rue de Prony, 75017 Paris, FRANCE
Telephone: 33-(0)1 44 15 18 88
Fax: 33-(0)1 42 67 09 87
Electronic mail: [email protected]
http://www.oie.int
All World Organisation for Animal Health (OIE) publications are protected by international copyright law. Extracts
may be copied, reproduced, translated, adapted or published in journals, documents, books, electronic media and
any other medium destined for the public, for information, educational or commercial purposes, provided prior
written permission has been granted by the OIE.
The designations and denominations employed and the presentation of the material in this publication do not
imply the expression of any opinion whatsoever on the part of the OIE concerning the legal status of any country,
territory, city or area or of its authorities, or concerning the delimitation of its frontiers and boundaries.
FOREWORD
The Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Terrestrial Manual) aims to
prevent and control animal diseases, including zoonoses, to contribute to the improvement of
animal health services world-wide and to allow safe international trade in animals and animal
products. The principal target readership is laboratories carrying out veterinary diagnostic tests and
surveillance, along with vaccine manufacturers and users, and regulatory authorities in Member
Countries. The main objective is to provide internationally agreed diagnostic laboratory methods
and requirements for the production and control of relevant vaccines and other biological products.
This ambitious task has required the cooperation of highly renowned animal health specialists from
many OIE Member Countries. The OIE, the World Organisation for Animal Health, received the
mandate from its Member Countries to undertake this task on a global level. The main activities of
the organisation, which was established in 1924, and in 2012 comprised 178 Member Countries,
are as follows:
1.
To ensure transparency in the global animal disease and zoonosis situation.
2.
To collect, analyse and disseminate scientific veterinary information on animal disease control
methods.
3.
To provide expertise and encourage international solidarity in the control of animal diseases.
4.
Within its mandate under the WTO (World Trade Organization) Agreement on Sanitary and
Phytosanitary Measures (SPS Agreement), to safeguard world trade by publishing health
standards for international trade in animals and animal products.
5.
To improve the legal framework and resources of national Veterinary Services.
6.
To provide a better guarantee of the safety of food of animal origin and to promote animal
welfare through a science-based approach.
The Terrestrial Manual, covering infectious and parasitic diseases of mammals, birds and bees,
was first published in 1989. Each successive edition has extended and updated the information
provided. This seventh edition includes over 50 updated chapters and guidelines (including a new
guideline on the application of biotechnology to the development of veterinary vaccines, and the
addition of epizootic haemorrhagic disease to the relevant chapter). This edition has a slightly
different structure from former editions: Part 1 contains ten introductory chapters that set general
standards for the management of veterinary diagnostic laboratories and vaccine production
facilities; Part 2 comprises chapters on OIE listed diseases and other diseases of importance to
international trade; Part 3 comprises four guidelines that have been developed on topics such as
biotechnology and antimicrobial susceptibility testing that are intended to give a brief introduction to
their subjects (they are to be regarded as background information rather than strict standards); and
Part 4 is the list of OIE Reference Centres at the time of publication (the list of OIE Reference
Centres is updated by the World Assembly of Delegates (of OIE Member Countries) each year; the
revised list is available on the OIE Web site).
As a companion volume to the Terrestrial Animal Health Code, the Terrestrial Manual sets
laboratory standards for all OIE listed diseases as well as several other diseases of global
importance. In particular it specifies (in blue font) those “Prescribed Tests” that are recommended
for use in health screening for international trade or movement of animals. The Terrestrial Manual
has become widely adopted as a key reference book for veterinary laboratories around the world.
Aquatic animal diseases are included in a separate Aquatic Manual.
The task of commissioning chapters and compiling the Terrestrial Manual was assigned to the OIE
Biological Standards Commission by the World Assembly of national Delegates. Manuscripts were
requested from specialists (usually the OIE designated experts at OIE Reference Laboratories) in
each of the diseases or the other topics covered. Occasionally, an ad hoc Group of experts was
convened tasked with updating or developing a chapter. After initial scrutiny by the Consultant
Technical Editor, the chapters were sent to scientific reviewers and to experts at OIE Reference
Laboratories. They were also circulated to all OIE Member Countries for review and comment. The
Biological Standards Commission, elected every 3 years by the World Assembly, and the
OIE Terrestrial Manual 2012
iii
Foreword
Consultant Technical Editor took all the resulting comments into consideration, often referring back
to the contributors for further help, before finalising the chapters. The final text has the approval of
the World Assembly.
A procedure for the official recognition of commercialised diagnostic tests, under the authority of the
Assembly, was finalised in September 2004. Data are submitted using a validation template that
was developed by the Biological Standards Commission. Submissions are evaluated by appointed
experts, who advise the Biological Standards Commission before the final opinion of the OIE World
Assembly is sought. All information on the submission of applications can be found on the OIE Web
site.
The Terrestrial Manual continues to expand and to extend its range of topics covered. It is our
sincere hope that it will grow in usefulness to veterinary diagnosticians and vaccine manufacturers
in all the OIE Member Countries. A new paper edition of the Terrestrial Manual is published every
4 years. It is important to note that annual updates to the Terrestrial Manual will be published on the
OIE website once approved by the World Assembly, so readers are advised to check there for the
latest information. This new version of the Terrestrial Manual is published in English and Spanish.
iv
Doctor Bernard Vallat
Director General, OIE
Professor Vincenzo Caporale
President, OIE Biological Standards Commission
September 2012
September 2012
OIE Terrestrial Manual 2012
ACKNOWLEDGEMENTS
I am most grateful to the many people whose combined efforts have gone into the preparation of
this Terrestrial Manual. In particular, I would like to express my thanks to:
Dr Bernard Vallat, Director General of the OIE from 2001 to the present, who gave his
encouragement and support to the project of preparing the new edition of this Terrestrial
Manual,
The Members of the OIE Standards Commission, Prof. Vincenzo Caporale, Dr Beverly
Schmitt, Dr Mehdi El Harrak, Dr Paul Townsend, Dr Alejandro Schudel and Dr Hualan Chen
who were responsible for commissioning chapters and, with the Consultant Technical Editor,
for editing all the contributions so as to finalise this edition of the Terrestrial Manual,
The contributors listed on pages xxii to xxxv who contributed their invaluable time and
expertise to write the chapters,
The expert advisers to the Biological Standards Commission’s meeting, Dr Adama Diallo and
Dr Peter Wright, the OIE Reference Laboratory experts and other reviewers who also gave
their time and expertise to scrutinising the chapters,
Those OIE Member Countries that submitted comments on the draft chapters that were
circulated to them. These were essential in making the Terrestrial Manual internationally
acceptable,
Ms Sara Linnane who, as Scientific Editor, organised this complex project and made major
contributions to the quality of the text,
Prof. Steven Edwards, Consultant Technical Editor of the Terrestrial Manual, who contributed
hugely to editing and harmonising the contents, but also in collating and incorporating Member
Country comments,
Members of both the OIE Scientific and Technical Department and the Publications
Department, for their assistance.
Dr Karin Schwabenbauer
President of the OIE World Assembly
September 2012
OIE Terrestrial Manual 2012
v
CONTENTS
VOLUME 2
Introduction (How to use this Terrestrial Manual) .....................................................
List of tests for International trade ............................................................................
Common abbreviations used in this Terrestrial Manual ............................................
Glossary of terms .....................................................................................................
Contributors ..............................................................................................................
PART 2
CONTINUED
OIE LISTED DISEASES AND OTHER DISEASES OF IMPORTANCE TO
INTERNATIONAL TRADE
SECTION 2.5.
EQUIDAE
Chapter 2.5.1.
Chapter 2.5.2.
Chapter 2.5.3.
Chapter 2.5.4.
Chapter 2.5.5.
Chapter 2.5.6.
Chapter 2.5.7.
Chapter 2.5.8.
Chapter 2.5.9.
Chapter 2.5.10.
Chapter 2.5.11.
Chapter 2.5.12.
Chapter 2.5.13.
African horse sickness .............................................................................................
Contagious equine metritis .......................................................................................
Dourine .....................................................................................................................
Epizootic lymphangitis ..............................................................................................
Equine encephalomyelitis (Eastern and Western) ....................................................
Equine infectious anaemia .......................................................................................
Equine influenza .......................................................................................................
Equine piroplasmosis ...............................................................................................
Equine rhinopneumonitis ..........................................................................................
Equine viral arteritis ..................................................................................................
Glanders ...................................................................................................................
Horse mange ............................................................................................................
Venezuelan equine encephalomyelitis .....................................................................
SECTION 2.6.
LEPORIDAE
Chapter 2.6.1.
Chapter 2.6.2.
Myxomatosis ............................................................................................................
Rabbit haemorrhagic disease ...................................................................................
SECTION 2.7.
CAPRINAE
Chapter 2.7.1.
Chapter 2.7.2.
Chapter 2.7.3/4.
Chapter 2.7.5.
Chapter 2.7.6.
Chapter 2.7.7.
Chapter 2.7.8.
Chapter 2.7.9.
Chapter 2.7.10
Chapter 2.7.11.
Chapter 2.7.12.
Chapter 2.7.13.
Chapter 2.7.14.
Border disease .........................................................................................................
Caprine and ovine brucellosis (excluding Brucella ovis) ...........................................
Caprine arthritis/encephalitis and Maedi-visna .........................................................
Contagious agalactia ................................................................................................
Contagious caprine pleuropneumonia ......................................................................
Enzootic abortion of ewes (ovine chlamydiosis) .......................................................
Nairobi sheep disease ..............................................................................................
Ovine epididymitis (Brucella ovis).............................................................................
Ovine pulmonary adenocarcinoma (adenomatosis) .................................................
Peste des petits ruminants .......................................................................................
Salmonellosis (S. abortusovis) .................................................................................
Scrapie .....................................................................................................................
Sheep pox and goat pox ..........................................................................................
OIE Terrestrial Manual 2012
ix
xi
xv
xvii
xxii
819
831
838
846
852
860
865
879
889
899
913
923
924
931
941
957
968
978
987
995
1008
1017
1018
1027
1032
1044
1045
1055
vii
Contents
SECTION 2.8.
SUIDAE
Chapter 2.8.1.
Chapter 2.8.2.
Chapter 2.8.3.
Chapter 2.8.4.
Chapter 2.8.5.
Chapter 2.8.6.
Chapter 2.8.7.
Chapter 2.8.8.
Chapter 2.8.9.
Chapter 2.8.10.
Chapter 2.8.11.
African swine fever ...................................................................................................
Atrophic rhinitis of swine ...........................................................................................
Classical swine fever (hog cholera) ..........................................................................
Nipah virus encephalitis ...........................................................................................
Porcine brucellosis ...................................................................................................
Porcine cysticercosis ................................................................................................
Porcine reproductive and respiratory syndrome .......................................................
Swine influenza ........................................................................................................
Swine vesicular disease ...........................................................................................
Teschovirus encephalomyelitis (previously enterovirus encephalomyelitis or
Teschen/Talfan disease) ..........................................................................................
Transmissible gastroenteritis ....................................................................................
SECTION 2.9.
OTHER DISEASES1
Chapter 2.9.1.
Chapter 2.9.2.
Chapter 2.9.3.
Chapter 2.9.4.
Chapter 2.9.5.
Chapter 2.9.6.
Chapter 2.9.7.
Chapter 2.9.8.
Chapter 2.9.9.
Chapter 2.9.10.
Chapter 2.9.11.
Chapter 2.9.12.
Bunyaviral diseases of animals (excluding Rift Valley fever)* ..................................
Camelpox .................................................................................................................
Campylobacter jejuni and Campylobacter coli..........................................................
Cryptosporidiosis ......................................................................................................
Cysticercosis* ...........................................................................................................
Hendra and Nipah virus diseases.............................................................................
Listeria monocytogenes. ..........................................................................................
Mange* .....................................................................................................................
Salmonellosis* ..........................................................................................................
Toxoplasmosis .........................................................................................................
Verocytotoxigenic Escherichia coli ...........................................................................
Zoonoses transmissible from non-human primates ..................................................
PART 3
GENERAL GUIDELINES
Guideline 3.1.
Guideline 3.2.
Guideline 3.3.
Guideline 3.4.
Laboratory methodologies for bacterial antimicrobial susceptibility testing ..............
Biotechnology in the diagnosis of infectious diseases ..............................................
The application of biotechnology to the development of veterinary vaccines ...........
The role of official bodies in the international regulation of veterinary biologicals ....
PART 4
OIE REFERENCE EXPERTS AND DISEASE INDEX
List of OIE Reference Laboratories (as of May 2012) ...............................................................................
Alphabetical list of diseases ......................................................................................................................
1
viii
1067
1080
1089
1105
1106
1113
1114
1127
1138
1145
1152
1163
1175
1183
1190
1214
1225
1239
1256
1268
1287
1297
1308
1311
1322
1338
1348
1363
1403
The diseases on this list that are marked with an asterisk are included in some individual species sections of the OIE List,
but these Terrestrial Manual chapters cover several species and thus give a broader description.
OIE Terrestrial Manual 2012
INTRODUCTION
(How to use this Terrestrial Manual)
•
Arrangement of the Terrestrial Manual
Part 1, the beginning of this Terrestrial Manual, contains ten introductory chapters that set general
standards for the management of veterinary diagnostic laboratories and vaccine facilities.
The main part of the Terrestrial Manual (Part 2) covers standards for diagnostic tests and vaccines
for specific diseases listed in the OIE Terrestrial Animal Health Code. The diseases are in
alphabetical order, subdivided by animal host group. OIE listed diseases are transmissible diseases
that have the potential for very serious and rapid spread, irrespective of national borders. They
have particularly serious socio-economic or public health consequences and are of major
importance in the international trade of animals and animal products.
Four of the diseases in Section 2.9 are included in some individual species sections, but these
chapters cover several host species and thus give a broader description. Some additional diseases
that may also be of importance to trade but that do not have a chapter in the Terrestrial Code are
also included in Section 2.9. This section also includes some important zoonotic infections
The contributors of all the chapters are listed on pages xxii–xxxv, but the final responsibility for the
content of the Terrestrial Manual lies with the World Assembly of the OIE.
There is an alphabetical index of the diseases at the end of Volume 2.
• Format of chapters
Each disease chapter includes a summary intended to provide information for veterinary officials
and other readers who need a general overview of the tests and vaccines available for the disease.
This is followed by a text giving greater detail for laboratory workers. In each disease chapter,
Part A gives a general introduction to the disease, Part B deals with laboratory diagnosis of the
disease, and Part C (where appropriate) with the requirements for vaccines or in vivo diagnostic
biologicals. The information concerning production and control of vaccines or diagnostics is given
as an example; it is not always necessary to follow these when there are scientifically justifiable
reasons for using alternative approaches. Bibliographic references that provide further information
are listed at the end of each chapter.
• Explanation of the tests described and of the table on pages xi–xiv
The table on pages xi–xiv lists diagnostic tests in two categories: ‘prescribed’ and ‘alternative’.
Prescribed tests are those that are required by the Terrestrial Animal Health Code for the testing of
animals before they are moved internationally. In the Terrestrial Manual these tests are printed in
blue. At present it is not possible to have prescribed tests for every listed disease. ‘Alternative tests’
are those that are suitable for the diagnosis of disease within a local setting, and can also be used
in the import/export of animals after bilateral agreement. There are often other tests described in
the chapters, which may be useful for specific purposes such as diagnosis or surveillance as
indicated in the chapters.
OIE Terrestrial Manual 2012
ix
Introduction (How to use this Terrestrial Manual)
• General guidelines
Four guidelines that have been developed on topics such as biotechnology and antimicrobial
susceptibility testing are included in Part 3 of this Terrestrial Manual. These are intended to give a
brief introduction to their subjects. They are to be regarded as background information rather than
standards.
• List of OIE Reference Laboratories
A list of OIE Reference Laboratories is given in Part 4 of this Terrestrial Manual. These laboratories
have been designated by the OIE as centres of excellence with expertise in their particular field.
They are able to provide advice to other laboratories on methodology. In some cases standard
strains of micro-organisms or reference reagents (e.g. antisera, antigens) can also be obtained
from the OIE Reference Laboratories.
The list of OIE Reference Laboratories will be updated by the World Assembly each year. The
revised list is available on the OIE Web site.
*
* *
x
OIE Terrestrial Manual 2012
LIST OF TESTS FOR INTERNATIONAL TRADE
The table below lists diagnostic tests in two categories: ‘prescribed’ and ‘alternative’. Prescribed tests
are required by the OIE Terrestrial Animal Health Code for the international movement of animals
and animal products and are considered optimal for determining the health status of animals. In the
Terrestrial Manual these tests are printed in blue. At present it is not possible to have prescribed
tests for every listed disease. Alternative tests are those that are suitable for the diagnosis of disease
within a local setting, and can also be used in the import/export of animals after bilateral agreement.
There are often other tests described in the chapters that may also be of some practical value in
local situations or that may still be under development.
Chapter No.
*
Disease name
2.1.1.
Anthrax
2.1.2.
Aujeszky’s disease
2.1.3.
Bluetongue
2.1.4.
Echinococcosis/Hydatidosis
2.1.5.
Foot and mouth disease
2.1.6.
Prescribed tests
Alternative tests
–
–
ELISA, VN
–
Agent id.,
ELISA, PCR
AGID, VN
–
–
ELISA*, VN
CF
Heartwater
–
ELISA, IFA
2.1.7.
Japanese encephalitis
–
–
2.1.8.
Leishmaniosis
–
Agent id.
2.1.9.
Leptospirosis
–
MAT
2.1.10.
New World screwworm (Cochliomyia
hominivorax) and Old World screwworm
(Chrysomya bezziana)
–
Agent id.
2.1.11.
Paratuberculosis (Johne’s disease)
–
DTH, ELISA
2.1.12.
Q fever
–
CF
2.1.13.
Rabies
ELISA, VN
–
2.1.14.
Rift Valley fever
VN
ELISA, HI
2.1.15.
Rinderpest
ELISA
VN
2.1.16.
Trichinellosis
Agent id.
ELISA
2.1.17.
Trypanosoma evansi infections
(including surra)
–
–
2.1.18.
Tularemia
–
Agent id.
2.1.19.
Vesicular stomatitis
CF, ELISA, VN
–
2.1.20.
West Nile fever
–
–
2.2.1.
Acarapisosis of honey bees
–
–
2.2.2.
American foulbrood of honey bees
–
–
Please refer to Terrestrial Manual chapters to verify which method is prescribed.
OIE Terrestrial Manual 2012
xi
List of tests for international trade
Chapter No.
xii
Disease name
Prescribed tests
Alternative tests
2.2.3.
European foulbrood of honey bees
–
–
2.2.4.
Nosemosis of bees
–
–
2.2.5.
Small hive beetle infestation
(Aethina tumida)
–
–
2.2.6.
Tropilaelaps infestation of honey bees
(Tropilaelaps spp.)
–
–
2.2.7
Varroosis of honey bees
–
–
2.3.1.
Avian chlamydiosis
–
–
2.3.2.
Avian infectious bronchitis
–
ELISA, HI, VN
2.3.3.
Avian infectious laryngotracheitis
–
AGID, ELISA, VN
2.3.4.
Avian influenza
Virus isolation
with pathogenicity
testing
AGID, HI
2.3.5.
Avian mycoplasmosis
(Mycoplasma gallisepticum, M. synoviae)
–
Agg., HI
2.3.6.
Avian tuberculosis
–
Agent id.,
Tuberculin test
2.3.7.
Duck virus enteritis
–
–
2.3.8.
Duck virus hepatitis
–
–
2.3.9.
Fowl cholera
–
–
2.3.10.
Fowl pox
–
–
2.3.11.
Fowl typhoid and Pullorum disease
–
Agent id., Agg.
2.3.12.
Infectious bursal disease
(Gumboro disease)
–
AGID, ELISA
2.3.13.
Marek’s disease
–
AGID
2.3.14.
Newcastle disease
Virus isolation
HI
2.3.15.
Turkey rhinotracheitis (avian
metapneumovirus)
–
–
2.4.1.
Bovine anaplasmosis
–
CAT, CF
2.4.2.
Bovine babesiosis
–
CF, ELISA, IFA
2.4.3.
Bovine brucellosis
BBAT, CF,
ELISA, FPA
–
2.4.4.
Bovine cysticercosis
–
Agent id.
2.4.5.
Bovine genital campylobacteriosis
Agent id.
–
2.4.6.
Bovine spongiform encephalopathy
–
–
2.4.7.
Bovine tuberculosis
Tuberculin test
Gamma interferon test
2.4.8.
Bovine viral diarrhoea
Agent id.
–
2.4.9.
Contagious bovine pleuropneumonia
CF, ELISA
–
2.4.10.
Dermatophilosis
–
–
2.4.11.
Enzootic bovine leukosis
AGID, ELISA
PCR
2.4.12.
Haemorrhagic septicaemia
–
Agent id.
2.4.13.
Infectious bovine rhinotracheitis/
infectious pustular vulvovaginitis
Agent id.
(semen only),
ELISA, PCR, VN
–
OIE Terrestrial Manual 2012
List of tests for international trade
Chapter No.
Disease name
Prescribed tests
Alternative tests
2.4.14.
Lumpy skin disease
–
VN
2.4.15.
Malignant catarrhal fever
–
IFA, PCR, VN
2.4.16.
Theileriosis
Agent id., IFA
–
2.4.17.
Trichomonosis
Agent id.
Mucus agg.
2.4.18.
Trypanosomosis (Tsetse-transmitted)
–
IFA
2.5.1.
African horse sickness
CF, ELISA
Agent id. (real-time
PCR), VN
2.5.2.
Contagious equine metritis
Agent id.
–
2.5.3.
Dourine
CF
ELISA, IFA
2.5.4.
Epizootic lymphangitis
–
–
2.5.5.
Equine encephalomyelitis
(Eastern and Western)
–
CF, HI, PRN
2.5.6.
Equine infectious anaemia
AGID
ELISA
2.5.7.
Equine influenza
–
HI
2.5.8.
Equine piroplasmosis
ELISA, IFA
CF
2.5.9.
Equine rhinopneumonitis
–
VN
2.5.10.
Equine viral arteritis
Agent id.
(semen only), VN
–
2.5.11.
Glanders
CF
–
2.5.12.
Horse mange
–
Agent id.
2.5.13.
Venezuelan equine encephalomyelitis
–
CF, HI, PRN
2.6.1.
Myxomatosis
–
AGID, CF, IFA
2.6.2.
Rabbit haemorrhagic disease
–
ELISA, HI
2.7.1.
Border disease
Agent id.
–
2.7.2.
Caprine and ovine brucellosis
(excluding Brucella ovis)
BBAT, CF,
ELISA, FPA
Brucellin test
Caprine arthritis/encephalitis & Maedi-visna
AGID, ELISA
–
–
–
CF
–
2.7.3/4.
2.7.5.
Contagious agalactia
2.7.6.
Contagious caprine pleuropneumonia
2.7.7.
Enzootic abortion of ewes
(ovine chlamydiosis)
–
CF
2.7.8.
Nairobi sheep disease
–
–
2.7.9.
Ovine epididymitis (Brucella ovis)
CF
ELISA
2.7.10.
Ovine pulmonary adenocarcinoma
(adenomatosis)
–
–
2.7.11.
Peste des petits ruminants
VN
ELISA
2.7.12.
Salmonellosis (S. abortusovis)
–
–
2.7.13.
Scrapie
–
–
2.7.14.
Sheep pox and goat pox
–
VN
2.8.1.
African swine fever
ELISA
IFA
2.8.2.
Atrophic rhinitis of swine
–
–
OIE Terrestrial Manual 2012
xiii
List of tests for international trade
Chapter No.
Disease name
Prescribed tests
2.8.3.
Classical swine fever (hog cholera)
2.8.4.
Nipah virus encephalitis
2.8.5.
Porcine brucellosis
2.8.6.
Alternative tests
ELISA,
FAVN, NPLA
–
–
–
BBAT, CFT,
ELISA, FPA
–
Porcine cysticercosis
–
–
2.8.7.
Porcine reproductive and
respiratory syndrome
–
ELISA, IFA, IPMA
2.8.8.
Swine influenza
–
–
2.8.9.
Swine vesicular disease
VN
ELISA
2.8.10.
Teschovirus encephalomyelitis (previously
enterovirus encephalomyelitis or
Teschen/Talfan disease)
–
VN
2.8.11.
Transmissible gastroenteritis
–
VN, ELISA
2.9.1.
Bunyaviral diseases of animals (excluding
Rift Valley fever)
–
–
2.9.2.
Camelpox
–
–
2.9.3.
Campylobacter jejuni and C. coli
–
–
2.9.4.
Cryptosporidiosis
–
–
2.9.5.
Cysticercosis
–
Agent id.
2.9.6.
Hendra and Nipah virus diseases
–
–
2.9.8.
Listeria monocytogenes
–
–
2.9.8.
Mange
–
Agent id.
2.9.9.
Salmonellosis
–
Agent id.
2.9.10.
Toxoplasmosis
–
–
2.9.11.
Verocytotoxigenic Escherichia coli
–
–
2.9.12.
Zoonoses transmissible from non-human
primates
–
–
Note: The tests prescribed by the Terrestrial Animal Health Code for the purposes of international trade are printed
in blue in this Terrestrial Manual.
Abbreviations
Agent id.
Agent identification
HI
Haemagglutination inhibition
Agg.
Agglutination test
IFA
Indirect fluorescent antibody
AGID
Agar gel immunodiffusion
IPMA
Immunoperoxidase monolayer assay
BBAT
Buffered Brucella antigen test
MAT
Microscopic agglutination test
CAT
Card agglutination test
NPLA
Neutralising peroxidase-linked assay
CF
Complement fixation
PCR
Polymerase chain reaction
DTH
Delayed-type hypersensitivity
PRN
Plaque reduction neutralisation
ELISA
Enzyme-linked immunosorbent assay
VN
Virus neutralisation
FAVN
Fluorescent antibody virus neutralisation
–
No test designated yet
FPA
Fluorescence polarisation assay
xiv
OIE Terrestrial Manual 2012
COMMON ABBREVIATIONS USED
IN THIS TERRESTRIAL MANUAL
ABTS
FBS
Fetal bovine serum
FITC
FLK
Fluorescein isothiocyanate
Fetal lamb kidney (cells)
FPA
g
GIT
Fluorescence polarisation assay
Relative centrifugal force
Growth inhibition test
BHK
BLP
BPAT
BSA
BSF
Buffered Brucella antigen test
Bovine fetal kidney (cells)
Beef extract-glucose-peptone-serum
(medium)
Baby hamster kidney (cell line)
Buffered lactose peptone
Buffered plate antigen test
Bovine serum albumin
Bovine serum factors
HA
HAD
HBSS
HEP
HEPES
CAM
CEF
CF
CFU
CIEP
CK
Chorioallantoic membrane
Chicken embryo fibroblast
Complement fixation (test)
Colony-forming unit
Counter immunoelectrophoresis
Calf kidney (cells)
HI
HRPO
IB
ICFTU
ICPI
ID50
Haemagglutination
Haemadsorption
Hanks’ balanced salt solution
High-egg-passage (virus)
N-2-hydroxyethylpiperazine, N-2ethanesulphonic acid (buffer)
Haemagglutination inhibition
Horseradish peroxidase
Immunoblot test
International complement fixation test unit
Intracerebral pathogenicity index
Median infectious dose
CNS
CPE
CPLM
Central nervous system
Cytopathic effect
Cysteine-peptone-liver infusion maltose
(medium)
Casein-sucrose-yeast (agar)
Diethyl pyrocarbonate
Diethylaminoethyl
Diethylpyrocarbonate
Dulbecco’s modified Eagle’s medium
Dimethyl sulphide
Delayed-type hypersensitivity
Ethylene diamine tetra-acetic acid
Ethylene glycol tetra-acetic acid
Egg-infective dose
Enzyme-linked immunosorbent assay
Evans’ modified Tobie’s medium
Earle’s yeast lactalbumin (balanced salt
solution)
Fluorescent antibody test
Fluorescent antibody virus neutralisation
IFA
IHA
IPMA
Indirect fluorescent antibody (test)
Indirect haemagglutination
Immunoperoxidase monolayer assay
IU
IVPI
LA
LD
LEP
LPS
MAb
MAT
MCS
MDBK
MDT
MEM
MHC
International units
Intravenous pathogenicity index
Latex agglutination
Lethal dose
Low egg passage (virus)
Lipopolysaccharide
Monoclonal antibody
Microscopic agglutination test
Master cell stock
Madin-Darby bovine kidney (cell line)
Mean death time
Minimal essential medium
Major histocompatibility complex
MLV
m.o.i.
Modified live virus (vaccine)
multiplicity of infection
AGID
ATCC1
BBAT
BFK
BGPS
CSY
DEPC
DEAE
DEPC
DMEM
DMSO
DTH
EDTA
EGTA
EID
ELISA
EMTM
EYL
FAT
FAVN
1
2,2’-azino-di-(3-ethyl-benzthiazoline)-6sulphonic acid
Agar gel immunodiffusion
American type culture collection
American Type Culture Collection, P.O. Box 1549, Manassas, Virginia 20108, United States of America.
OIE Terrestrial Manual 2012
xv
Common abbreviations used in this Terrestrial Manual
MSV
NI
OGP
OPD
OPG
Master seed virus
Neutralisation index
1-octyl-beta-D-glucopyranoside (buffer)
Orthophenyldiamine (chromogen)
Oxalase-phenol-glycerin (preservative
solution)
Open reading frame
RBC
RFLP
RK
RPM
RSA
Red blood cell
Restriction fragment length polymorphism
Rabbit kidney
Revolutions per minute
Rapid serum agglutination
RT-PCR
SAT
SDS
PAS
PBS
PCR
PD
Polyacrylamide gel electrophoresis
Peroxidase–antiperoxidase (staining
procedure)
Periodic acid-Schiff (reaction)
Phosphate buffered saline
Polymerase chain reaction
Protective dose
Reverse-transcription polymerase chain
reaction
Serum agglutination test
Sodium dodecyl sulphate
SPF
SPG
SRBC
TCID50
Specific pathogen free
Sucrose phosphate glutamic acid
Sheep red blood cells
Median tissue culture infective dose
PFGE
PFU
PHA
PPD
PPLO
PRN
PSG
Pulsed field gel electrophoresis
Plaque-forming unit
Passive haemagglutination (test)
Purified protein derivative
Pleuropneumonia-like organisms
Plaque reduction neutralisation
Phosphate buffered saline glucose
TMB
TSI
VB
VBS
Vero
VN
Tetramethyl benzidine
Triple sugar iron (medium)
Veronal buffer
Veronal buffered saline
African green monkey kidney (cells)
Virus neutralisation
ORF
PAGE
PAP
*
* *
xvi
OIE Terrestrial Manual 2012
GLOSSARY OF TERMS
The definitions given below have been selected and restricted to those that are likely to be useful to
users of this OIE Terrestrial Manual.
•
Absorbance/optical density
Absorbance and optical density are terms used to indicate the strength of reaction. A spectrophotometer is used
to measure the amount of light of a specific wave length that a sample absorbs and the absorbance is
proportional to the amount of a particular analyte present.
•
Accuracy
Nearness of a test value to the expected value for a reference standard reagent of known activity or titre.
•
Assay
Synonymous with test or test method, e.g. enzyme immunoassay, complement fixation test or polymerase chain
reaction tests.
•
Batch
All vaccine or other reagent, such as antigen or antisera, derived from the same homogeneous bulk and identified
by a unique code number.
•
Cell line
A stably transformed line of cells that has a high capacity for multiplication in vitro.
•
Centrifugation
Throughout the text, the rate of centrifugation has been expressed as the Relative Centrifugal Force, denoted by
‘g’. The formula is:
(RPM × 0.10472)2
× Radius (cm) = g
980
where RPM is the rotor speed in revolutions per minute, and where Radius (cm) is the radius of the rotor arm, to
the bottom of the tube, in centimetres.
It may be necessary to calculate the RPM required to achieve a given value of g, with a particular rotor. The
formula is:
RPM =
√g × 980 /Radius (cm)
0.10472
•
Cross-reaction
See ’False-positive reaction’.
•
Cut-off/threshold
Test result value selected for distinguishing between negative and positive results; may include indeterminate or
suspicious zone.
OIE Terrestrial Manual 2012
xvii
Glossary of terms
•
Dilutions
Where dilutions are given for making up liquid reagents, they are expressed as, for example, 1 in 4 or 1/4,
meaning one part added to three parts, i.e. a 25% solution of A in B.
•
v/v – This is volume to volume (two liquids).
•
w/v – This is weight to volume (solid added to a liquid).
•
Dilutions used in virus neutralisation tests
There are two different conventions used in expressing the dilution used in virus neutralisation (VN) tests. In
Europe, it is customary to express the dilution before the addition of the antigen, but in the United States of
America and elsewhere, it is usual to express dilutions after the addition of antigen.
These alternative conventions are expressed in the Terrestrial Manual as ‘initial dilution’ or ‘final dilution’,
respectively.
•
Efficacy
Specific ability of the biological product to produce the result for which it is offered when used under the
conditions recommended by the manufacturer.
•
Equivalency testing
Determination of certain assay performance characteristics of new and/or different test methods by means of an
interlaboratory comparison to a standard test method; implied in this definition is that participating laboratories are
using their own test methods, reagents and controls and that results are expressed qualitatively.
•
False-negative reaction
Negative reactivity in an assay of a test sample obtained from an animal exposed to or infected with the organism
in question, may be due to lack of analytical sensitivity, restricted analytical specificity or analyte degradation,
decreases diagnostic sensitivity.
•
False-positive reaction
Positive reactivity in an assay that is not attributable to exposure to or infection with the organism in question,
maybe due to immunological cross-reactivity, cross-contamination of the test sample or non-specific reactions,
decreases diagnostic specificity.
•
Final product (lot)
All sealed final containers that have been filled from the same homogenous batch of vaccine in one working
session, freeze-dried together in one continuous operation (if applicable), sealed in one working session, and
identified by a unique code number.
•
Harmonisation
The result of an agreement between laboratories to calibrate similar test methods, adjust diagnostic thresholds
and express test data in such a manner as to allow uniform interpretation of results between laboratories.
•
Incidence
Estimate of the rate of new infections in a susceptible population over a defined period of time; not to be confused
with prevalence.
•
In-house checks
All quality assurance activities within a laboratory directly related to the monitoring, validation, and maintenance of
assay performance and technical proficiency.
•
In-process control
Test procedures carried out during manufacture of a biological product to ensure that the product will comply with
the agreed quality standards.
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OIE Terrestrial Manual 2012
Glossary of terms
•
Inter-laboratory comparison (ring test)
Any evaluation of assay performance and/or laboratory competence in the testing of defined samples by two or
more laboratories; one laboratory may act as the reference in defining test sample attributes.
•
Master cell (line, seed, stock)
Collection of aliquots of cells of defined passage level, for use in the preparation or testing of a biological product,
distributed into containers in a single operation, processed together and stored in such a manner as to ensure
uniformity and stability and to prevent contamination.
•
Master seed (agent, strain)
Collection of aliquots of an organism at a specific passage level, from which all other seed passages are derived,
which are obtained from a single bulk, distributed into containers in a single operation and processed together
and stored in such a manner as to ensure uniformity and stability and to prevent contamination.
•
Performance characteristic
An attribute of a test method that may include analytical sensitivity and specificity, accuracy and precision,
diagnostic sensitivity and specificity and/or repeatability and reproducibility.
•
Potency
Relative strength of a biological product as determined by appropriate test methods. (Initially the potency is
measured using an efficacy test in animals. Later this may be correlated with tests of antigen content, or antibody
response, for routine batch potency tests.)
•
Precision
The degree of dispersion of results for a repeatedly tested sample expressed by statistical methods such as
standard deviation or confidence limits.
•
Predictive value (negative)
The probability that an animal is free from exposure or infection given that it tests negative; predictive values are a
function of the DSe (diagnostic sensitivity) and DSp (diagnostic specificity) of the diagnostic assay and the
prevalence of infection.
•
Predictive value (positive)
The probability that an animal has been exposed or infected given that it tests positive; predictive values are a
function of the DSe and DSp of the diagnostic assay and the prevalence of infection.
•
Prevalence
Estimate of the proportion of infected animals in a population at one given point in time; not to be confused with
incidence.
•
Primary cells
A pool of original cells derived from normal tissue up to and including the tenth subculture.
•
Production seed
An organism at a specified passage level that is used without further propagation for initiating preparation of a
production bulk.
•
Proficiency testing
One measure of laboratory competence derived by means of an interlaboratory comparison; implied in this
definition is that participating laboratories are using the same test methods, reagents and controls and that results
are expressed qualitatively.
OIE Terrestrial Manual 2012
xix
Glossary of terms
•
Purity
Quality of a biological product prepared to a final form and:
a)
Relatively free from any extraneous microorganisms and extraneous material (organic or inorganic) as
determined by test methods appropriate to the product; and
b)
Free from extraneous microorganisms or material which could adversely affect the safety, potency or
efficacy of the product.
•
Reference animal
Any animal for which the infection status can be defined in unequivocal terms; may include diseased, infected,
vaccinated, immunised or naïve animals.
•
Reference Laboratory
Laboratory of recognised scientific and diagnostic expertise for a particular animal disease and/or testing
methodology; includes capability for characterising and assigning values to reference reagents and samples.
•
Repeatability
Level of agreement between replicates of a sample both within and between runs of the same test method in a
given laboratory.
•
Reproducibility
Ability of a test method to provide consistent results when applied to aliquots of the same sample tested by the
same method in different laboratories.
•
Room temperature
The term ‘room temperature’ is intended to imply the temperature of a comfortable working environment. Precise
limits for this cannot be set, but guiding figures are 18–25°C. Where a test specifies room temperature, this
should be achieved, with air conditioning if necessary; otherwise the test parameters may be affected.
•
Safety
Freedom from properties causing undue local or systemic reactions when used as recommended or suggested by
the manufacturer and without known hazard to in-contact animals, humans and the environment.
•
Sample
Material that is derived from a specimen and used for testing purposes.
•
Sensitivity (analytical)
Synonymous with ‘Limit of Detection’, smallest detectable amount of analyte that can be measured with a defined
certainty; analyte may include antibodies, antigens, nucleic acids or live organisms.
•
Sensitivity (diagnostic)
Proportion of known infected reference animals that test positive in the assay; infected animals that test negative
are considered to have false-negative results.
•
Sensitivity (relative)
Proportion of reference animals defined as positive by one or a combination of test methods that also test positive
in the assay being compared.
•
Specific pathogen free (SPF)
Animals that have been shown by the use of appropriate tests to be free from specified pathogenic
microorganisms, and also refers to eggs derived from SPF birds.
xx
OIE Terrestrial Manual 2012
Glossary of terms
•
Specificity (analytical)
Degree to which the assay distinguishes between the target analyte and other components in the sample matrix;
the higher the analytical specificity, the lower the level of false-positives.
•
Specificity (diagnostic)
Proportion of known uninfected reference animals that test negative in the assay; uninfected reference animals
that test positive are considered to have false-positive results.
•
Specificity (relative)
Proportion of reference animals defined as negative by one or a combination of test methods that also test
negative in the assay being compared.
•
Specimen
Material submitted for testing.
•
Standard Reagents
•
International Standard Reagents
Standard reagents by which all other reagents and assays are calibrated; prepared and distributed by an
International Reference Laboratory.
•
National Standard Reagents
Standard reagents calibrated by comparison with International Standard Reagents; prepared and distributed
by a National Reference Laboratory.
•
Working Standards (reagents)
Standard reagents calibrated by comparison with the National Standard Reagent, or, in the absence of a
National Standard Reagent, calibrated against a well-characterised in-house standard reagent; included in
routine diagnostic tests as a control and/or for normalisation of test results.
•
Sterility
Freedom from viable contaminating microorganisms, as demonstrated by approved and appropriate tests.
•
Test method
Specified technical procedure for detection of an analyte (synonymous with assay).
•
Tests
•
Prescribed
Test methods that are required by the OIE Terrestrial Animal Health Code for the international movement of
animals and animal products and that are considered optimal for determining the health status of animals.
•
Alternative
Test methods considered in this Terrestrial Manual to be suitable for the diagnosis of disease in a local
situation, and that can also be used for import/ export by bilateral agreement.
•
Screening
Tests of high diagnostic sensitivity suitable for large-scale application.
•
Confirmatory
Test methods of high diagnostic specificity that are used to confirm results, usually positive results, derived
from other test methods
•
Working seed
Organism at a passage level between master seed and production seed.
OIE Terrestrial Manual 2012
xxi
CONTRIBUTORS
C O N T R IB U T O R S A N D P R O F E S S I O N A L A D D R E S S A T T I ME O F W R I T IN G
The chapters in the Terrestrial Manual are prepared by invited contributors (OIE Reference Experts,
where possible). In accordance with OIE standard procedure, all chapters are circulated to OIE
Member Countries and to other experts in the disease for comment. The OIE Biological Standards
Commission and the Consultant Editor then modify the text to take account of comments received,
and the text is circulated a second time as the final version that will be presented for adoption by
the World Assembly of Delegates to the OIE at the General Session in May of each year. The
Terrestrial Manual is thus deemed to be an OIE Standard Text that has come into being by
international agreement. For this reason, the names of the contributors are not shown on individual
chapters but are listed below. The Biological Standards Commission greatly appreciates the work of
the following contributors (address at the time of writing):
1.1.1. Collection and shipment of
diagnostic specimens
Dr J.E. Pearson (retired)
4016 Phoenix St., Ames, Iowa 50014, USA.
1.1.2. Sampling of diagnostic specimens
Chapter under study
1.1.3. Biosafety and biosecurity in the veterinary
microbiology laboratory and animal facilities
Dr B. Schmitt
National Veterinary Services Laboratories,
Diagnostic Virology Laboratory, P.O. Box 844,
Ames, IA 50010, USA.
Dr P. Le Blanc Smith
CSIRO Livestock Industries, Australian Animal
Health Laboratory (AAHL), Private Bag 24,
Geelong, Victoria 3220,
Australia.
1.1.4. Quality management in veterinary testing
laboratories
Dr R.A. Williams,
AHVLA-Penrith, Merrythought, Calthwaite,
Penrith, Cumbria CA11 9RR, UK.
1.1.5. Principles of validation of diagnostic assays for
infectious diseases1
Dr R.H. Jacobson (retired)
27801 Skyridge Drive, Eugene, Oregon 97405,
USA.
Dr P. Wright (retired)
Aquatic Animal Health, Fisheries and Oceans
Canada, 343 University Avenue, Moncton,
New Brunswick, E1C 9B6, Canada.
1
xxii
This chapter was updated by consensus of the OIE ad hoc Group on Validation of Diagnostic Assays.
OIE Terrestrial Manual 2012
Contributors
1.1.6. Principles of veterinary vaccine production
Dr B. Rippke
Center for Veterinary Biologics, USDA, Animal
and Plant Health Inspection Service, Veterinary
Services, Suite 104, 510 South 17th Street, Ames,
IA 50010, USA.
Dr D.J.K. Mackay
European Medicines Agency, Veterinary
Medicines and Inspections, 7 Westferry Circus,
Canary Wharf, London E14 4HB, UK.
Dr M. Lombard
International Association for Biologicals (IABs),
22 Rue Crillon, 69006 Lyon, France.
1.1.7. Tests for sterility and freedom from
contamination of biological materials
Dr L. Elsken
USDA, APHIS, Center for Veterinary Biologics,
Suite 104, 510 South 17th Street, Ames, Iowa
50010, USA.
1.1.8. Minimum requirements for vaccine production
facilities
Chapter under study
1.1.9. Quality control of vaccines
Chapter under study
1.1.10. International standards for vaccine banks
OIE ad hoc Group on Antigen and Vaccine Banks
for Foot and Mouth Disease
2.1.1. Anthrax
Dr E. Golsteyn-Thomas
Canadian Food Inspection Agency, Lethbridge
Laboratory, P.O. Box 640, Township Road 9-1
Lethbridge, Alberta T1J 3Z4, Canada.
Dr G. Harvey
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
2.1.2. Aujeszky’s disease
Dr A. Jestin & Dr M.F. Le Potier
ANSES Ploufragan, Laboratoire d’études et de
recherches avicoles et porcines, Zoopôle des
Côtes d’Armor-Les Croix, BP 53, 22440
Ploufragan, France.
2.1.3. Bluetongue
Dr P. Daniels
Australian Animal Health Laboratory, CSIRO
Livestock Industries, 5 Portarlington Road,
Geelong, Victoria 3220, Australia.
Dr C.A.L. Oura (formerly)
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Surrey GU24 0NF, UK.
2
2.1.4. Echinococcosis/Hydatidosis
Dr M. Kamiya
Laboratory of Environmental Zoology, Department
of Biosphere and Environmental Sciences,
Faculty of Environmental Systems, Rakuno
Gakuen University, Midori-machi 582,Ebetsu 0698501, Hokkaido, Japan.
2.1.5. Foot and mouth disease2
Dr D.J. Paton, Dr P.V. Barnett & Dr N.P. Ferris
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Surrey GU24 0NF, UK.
This chapter was updated by consensus of the OIE ad hoc Group on Vaccine Quality related to Foot and Mouth Disease.
OIE Terrestrial Manual 2012
xxiii
Contributors
2.1.6. Heartwater
Dr D. Martinez
CIRAD-EMVT, Campus International de
Baillarguet - TA30/G, 34398 Montpellier Cedex 5,
France.
Dr N. Vachiéry
CIRAD-EMVT, Domaine de Duclos, Prise d’Eau,
97170 Petit-Bourg, Guadeloupe.
Prof. F. Jongejan
Department of Parasitology & Tropical Veterinary
Medicine, Faculty of Veterinary Medicine, Utrecht
University, P.O. Box 80.165, 3508 TD Utrecht,
The Netherlands
AND
Department of Veterinary Tropical Diseases,
Faculty of Veterinary Science, University of
Pretoria, Onderstepoort 0110, South Africa.
2.1.7. Japanese encephalitis
Dr T. Kondo
Epizootic Research Center, Equine Research
Institute, Japan Racing Association, 1400-4
Shiba, Shimotsuke, Tochigi 329-0412, Japan.
2.1.8. Leishmaniosis
Dr L. Gradoni & Dr M. Gramiccia
Dipartimento di Malattie Infettive, Parassitarie ed
Immunomediate, Istituto Superiore di Sanità, Viale
Regina Elena 299, I-00161 Rome, Italy.
2.1.9. Leptospirosis
Prof. C.A. Bolin
Diagnostic Center for Population & Animal Health,
College of Veterinary Medicine, Michigan State
University, 4125 Beaumont Rd, Lansing, Michigan
48910, USA.
2.1.10. New World screwworm (Cochliomyia
hominivorax) and Old World screwworm
(Chrysomya bezziana)
Dr M.J.R. Hall
Department of Entomology, The Natural History
Museum, Cromwell Road, London SW7 5BD, UK.
2.1.11. Paratuberculosis (Johne’s disease)
Dr J. Gwozdz
Department of Primary Industries, Victoria,
475 Mickleham Road, Attwood, VIC 3049,
Australia.
2.1.12. Q fever
Dr E. Rousset, Dr K. Sidi-Boumedine &
Dr R. Thiery
Anses Sophia Antipolis, Laboratoire d’Études et
de Recherches sur les Petits Ruminants et les
Abeilles (LERPRA), Les Templiers, 105 route des
Chappes, BP 111, 06902 Sophia Antipolis Cedex,
France.
xxiv
OIE Terrestrial Manual 2012
Contributors
2.1.13. Rabies
Dr A. Fooks & Dr D. Horton
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
Dr T. Müller & Dr C. Freuling
Institute for Epidemiology, Friedrich-Loeffler
Institut, Federal Research Institute for Animal
Health, Seestr. 55, D-16868 Wusterhausen/Dosse
Germany.
Dr Charles Rupprecht
Centers for Disease Control and Prevention,
National Center for Emerging and Zoonotic
Infectious Diseases, Division of High
Consequence Pathogens and Pathology, Poxvirus
and Rabies Branch, 1600 Clifton Rd., MS G33
Atlanta, GA, 30333, USA.
2.1.14. Rift Valley fever
Dr G.H. Gerdes (retired)
Onderstepoort Veterinary Institute, Private Bag
X05, Onderstepoort 0110, South Africa.
2.1.15. Rinderpest
Dr W.P. Taylor
16 Mill Road, Angmering, Littlehampton, West
Sussex BN16 4HT, UK.
Dr P. Roeder
Hollyhedge Cottage, Spats Lane, Headley Down,
Bordon, Hampshire GU35 8SY, UK.
3
2.1.16. Trichinellosis
Dr A. Gajadhar & Dr L. Forbes
Canadian Food Inspection Agency, Centre for
Foodborne & Animal Parasitology, 116 Veterinary
Road, Saskatoon, Saskatchewan S7N 2R3
Canada.
2.1.17. Trypanosoma evansi infection (surra)3
Dr M. Desquesnes
UMR177-Intertryp (CIRAD-IRD), CIRAD-bios,
Campus international de Baillarguet, TA A-17 / G,
34398 Montpellier Cedex 5, France.
2.1.18. Tularemia
Dr T. Mörner (retired)
Department of Pathology and Wildlife Diseases,
Swedish National Veterinary Institute, Sweden.
2.1.19. Vesicular stomatitis
Dr S.L. Swenson
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
2.1.20. West Nile fever
Dr E.N. Ostlund
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
United States of America.
2.2.1. Acarapisosis of honey bees
Dr W. Ritter
CVUA-Freiberg, FB: Bienen (bee team), Am
Moosweiher 2, D79018 Freiberg, Germany.
2.2.2. American foulbrood of honey bees
2.2.3. European foulbrood of honey bees
Dr D.C. de Graaf
Laboratory of Zoophysiology, University of Ghent,
K.L. Ledeganckstraat 35, B-9000 Ghent, Belgium.
This chapter was updated by consensus of the OIE ad hoc Group on Diagnostic Tests for Trypanosomoses.
OIE Terrestrial Manual 2012
xxv
Contributors
2.2.4. Nosemosis of honey bees
Dr W. Ritter
CVUA-Freiberg, FB: Bienen (bee team), Am
Moosweiher 2, D79018 Freiberg, Germany.
2.2.5. Small hive beetle infestation (Aethina tumida)
Dr W. Ritter
CVUA-Freiberg, FB: Bienen (bee team), Am
Moosweiher 2, D79018 Freiberg, Germany.
Dr P. Neumann
Swiss Bee Research Centre, Agroscope
Liebefeld-Posieux, Research Station ALP,
Schwarzenburgstrasse 161, CH-3003 Bern,
Switzerland.
Dr J.D. Ellis
Department of Entomology, The University of
Georgia, Athens, GA 30602, USA.
2.2.6. Tropilaelaps infestation of honey bees
(Tropilaelaps spp.)
2.2.7. Varroosis of honey bees
Dr W. Ritter
CVUA-Freiberg, FB: Bienen (bee team), Am
Moosweiher 2, D79018 Freiberg, Germany.
2.3.1. Avian chlamydiosis
Dr K. Sachse
Friedrich-Loeffler-Institut, Federal Research
Institute for Animal Health, Institute of Molecular
Pathogenesis, Naumburger Str. 96a, 07743 Jena,
Germany.
2.3.2. Avian infectious bronchitis
Dr J. Gelb
Dept of Animal and Food Sciences and the Avian
Biosciences Center, University of Delaware,
531 South College Avenue, Newark, Delaware
19716-2150, USA.
2.3.3. Avian infectious laryngotracheitis
Dr R.C. Jones
Department of Veterinary Pathology, University of
Liverpool, Jordan Building, Veterinary Field
Station, ‘Leahurst’, Neston, South Wirral
CH64 7TE, UK.
2.3.4. Avian influenza
Dr D. Swayne
Southeast Poultry Research Laboratory,
934 College Station Road, Athens, Georgia 30605
USA.
2.3.5. Avian mycoplasmosis
(Mycoplasma gallisepticum, M. synoviae)
Dr S.H. Kleven
University of Georgia, Poultry Diagnostic and
Research Center, 953 College Stantion Road,
Athens, Georgia 30602-4875, USA.
Dr J.M. Bradbury
University of Liverpool, Department of Veterinary
Pathology, Veterinary Teaching Hospital,
Leahurst, Neston H64 7TE, UK.
2.3.6. Avian tuberculosis
xxvi
Dr D.V. Cousins (formerly)
Australian Reference Laboratory for Bovine
Tuberculosis, Western Australia Dept of
Agriculture and Food, Locked Bag N° 4, Bentley
Delivery Centre, Bentley WA 6983, Australia.
OIE Terrestrial Manual 2012
Contributors
2.3.7. Duck virus enteritis
2.3.8. Duck virus hepatitis
Dr P.R. Woolcock
California Animal Health and Food Safety
Laboratory System, University of California, Davis
West Health Sciences Drive, P.O. Box 1770,
Davis, California 95617, USA.
2.3.9. Fowl cholera
Dr R. Kunkle
National Animal Disease Center, P.O. Box 70,
Ames, Iowa 50010, USA.
Dr M.A. Wilson
National Animal Disease Center, 1800 N. Dayton
Avenue, Ames, Iowa 50010, USA.
2.3.10. Fowl pox
Dr D.N. Tripathy
University of Illinois at Urbana-Champaign,
College of Veterinary Medicine, Department of
Veterinary Pathbiology, 2001 South Lincoln
Avenue, Urbana, Illinois 61802, USA.
2.3.11. Fowl typhoid and Pullorum disease
Dr R. Davies
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
2.3.12. Infectious bursal disease (Gumboro disease)
Dr N. Eterradossi
Anses, Laboratoire de Ploufragan-Plouzané,
Laboratoire d'études et de recherches avicoles,
porcines et piscicoles, B.P. 53, 22440 PloufraganPlouzané, France.
2.3.13. Marek’s disease
Dr Venugopal Nair
Institute for Animal Health, Compton, Berkshire
RG20 7NN, UK.
2.3.14. Newcastle disease
Dr C.L. Afonso & Dr P.J. Miller
SEPRL, USDA, ARS, SAA, 934 College Station
Road, Athens, GA 30605, USA.
Dr Ch. Grund
Friedrich-Loeffler-Institute, Federal Research
Institute for Animal Health, Institute of Diagnostic
Virology, Südufer 10, D-17493 Greifswald, Insel
Riems, Germany.
Dr G. Koch & Dr B. Peeters
Central Veterinary Institute of Wageningen
University and Research Centre, Department of
Virology, P.O. Box 65, NL-8200 AB Lelystad, The
Netherlands.
Dr Paul W. Selleck
CSIRO, Australian Animal Health Laboratory
(AAHL), 5 Portarlington Road, Private Bag 24,
Geelong 3220, Victoria, Australia.
Dr G.B. Srinivas
Bacteriology Policy, Evaluation, and Licensing,
USDA/APHIS/VS/CVB, 1920 Dayton Avenue,
P.O. Box 844 Ames, Iowa 50010, USA.
OIE Terrestrial Manual 2012
xxvii
Contributors
2.3.15. Turkey rhinotracheitis (avian
metapneumovirus)
Dr J. Pedersen
Avian Section, Diagnostic Virology Laboratory,
National Veterinary Services Laboratories,
1920 Dayton Avenue, Ames, IA 50010, USA.
Dr N. Eterradossi
Anses, Laboratoire de Ploufragan-Plouzané,
Laboratoire d'études et de recherches avicoles,
porcines et piscicoles, B.P. 53, 22440 PloufraganPlouzané, France.
2.4.1. Bovine anaplasmosis
2.4.2. Bovine babesiosis
Prof. T.F. McElwain
Animal Health Research Center, College of
Veterinary Medicine, 155N Bustad Hall, P.O. Box
647034, Pullman, WA 99164-7034, USA.
Dr P.J. Rolls, Dr R.E. Bock, Dr A.J. de Vos &
Dr S.J. Waldron
Tick Fever Centre, Biosecurity Queensland
280 Grindle Road, Wacol, Queensland 4054,
Australia.
Dr I.E. Echaide
Instituto Nacional de Tecnología Agropecuaria,
Estación Experimental Agropecuaria Rafaela,
CC22, CP 2300 Rafaela (Santa Fe), Argentina.
2.4.3. Bovine brucellosis4
Dr K. Nielsen (retired)
Canadian Food Inspection Agency, Animal
Diseases Research Institute, P.O. Box 11300,
Station H, Nepean, Ontario K2H 8P9, Canada.
Dr D.R. Ewalt
Pathobiology Laboratory, National Veterinary
Services Laboratories, 1800 Dayton Road, Ames,
Iowa 50010, USA.
2.4.4. Bovine cysticercosis
See chapter 2.9.5.
2.4.5. Bovine genital campylobacteriosis
Prof. J.A. Wagenaar
Utrecht University, Faculty of Veterinary Medicine,
P.O. Box 80.165, 3508 TD Utrecht, The
Netherlands.
Dr M.A.P. Van Bergen
Central Veterinary Institute of Wageningen UR,
P.O. Box 65, 8200 AB Lelystad, The Netherlands.
4
2.4.6. Bovine spongiform encephalopathy
Dr M.M. Simmons, Mr M.J. Stack, Dr T. Konold &
Dr K. Webster
AHVLA Weybridge, New Haw, Addlestone,
Surrey KT15 3NB, UK.
2.4.7. Bovine tuberculosis
Dr D.V. Cousins (formerly)
Australian Reference Laboratory for Bovine
Tuberculosis, Western Australia Dept of
Agriculture and Food, Locked Bag N° 4, Bentley
Delivery Centre, Bentley WA 6983, Australia.
2.4.8. Bovine viral diarrhoea
Dr T. Drew
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
All four brucellosis chapters were updated by consensus of the OIE ad hoc Group on Brucellosis.
xxviii
OIE Terrestrial Manual 2012
Contributors
2.4.9. Contagious bovine pleuropneumonia
Dr F. Thiaucourt
CIRAD-EMVT, Campus international de
Baillarguet, Montferriez-sur-Lez, B.P. 5035, 34032
Montpellier Cedex 1, France.
2.4.10. Dermatophilosis
Dr D. Martinez
CIRAD, Campus International de Baillarguet – TAA15 / G, 34398 Montpellier Cedex 5, France.
2.4.11. Enzootic bovine leukosis
Dr T.W. Vahlenkamp
Institute of Virology, Centre for Infectious
Diseases, Faculty of Veterinary Medicine, Leipzig
University, An den Tierkliniken 29, 04103 Leipzig,
Germany.
2.4.12. Haemorrhagic septicaemia
Dr V.P. Singh
Indian Veterinary Research Institute, Izatnagar
243122 U.P., India.
2.4.13. Infectious bovine rhinotracheitis/
infectious pustular vulvovaginitis
Dr M. Beer
Institute of Diagnostic Virology, Friedrich-LoefflerInstitut, Südufer 10, D-17493 Greifswald-Insel
Riems, Germany.
2.4.14. Lumpy skin disease
Dr E. Tuppurainen
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Woking, Surrey GU24 0NF,
UK.
2.4.15. Malignant catarrhal fever
Dr H.W. Reid
Moredun Research Institute, International
Research Centre, Pentlands Science Park, Bush
Loan, Penicuik EH26 0PZ, Scotland, UK.
2.4.16. Theileriosis
Prof. E. Pipano
Koret School of Veterinary Medicine, The Hebrew
University of Jerusalem, P.O. Box 12 Rehovot,
Israel.
Dr S. Morzaria
FAO Regional Office for Asia and the Pacific,
39 Phra Athit Road, Bangkok 10200, Thailand.
Dr P. Spooner
International Livestock Research Institute,
Naivasha Road, Nairobi 00100, Kenya.
2.4.17. Trichomonosis
Dr A.A. Gajadhar
Centre for Food-borne and Animal Parasitology,
Canadian Food Inspection Agency,
116 Veterinary Road, Saskatoon,
Saskatchewan S7N 2R3, Canada.
Dr S. Parker
Large Animal Clinical Sciences, Western College
of Veterinary Medicine,52 Campus Drive,
Saskatoon, Saskatchewan S7N 5B4, Canada.
2.4.18. Trypanosomosis (Tsetse-transmitted)
OIE Terrestrial Manual 2012
Dr M. Desquesnes
UMR177-Intertryp (CIRAD-IRD), CIRAD-bios,
Campus international de Baillarguet, TA A-17 / G,
34398 Montpellier Cedex 5, France.
xxix
Contributors
2.5.1. African horse sickness
Prof. J.M. Sánchez-Vizcaíno
Centro de Vigilancia Sanitaria Veterinaria
(VISAVET), Facultad de Veterinaria, HCV Planta
sótano, Universidad Complutense de Madrid
(UCM), Avda Puerta de Hierro s/n, 28040 Madrid
Spain.
Dr C. Gómez-Tejedor Ortiz
Ministerio de Agricultura, Pesca y Alimentación
Laboratorio Central de Veterinaria, LCV-Algete,
Ctra. Algete Km 8, 28110 Algete (Madrid), Spain.
2.5.2. Contagious equine metritis
Mr P. Todd
AHVLA Bury St Edmunds, Rougham Hill, Bury St
Edmunds, Suffolk IP44, 2RX, UK
Dr M.M. Erdman
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
H.J. Roest
Central Veterinary Institute of Wageningen UR,
Department of Bacteriology, P.O. Box 65, 8200
AB Lelystad, The Netherlands.
2.5.3. Dourine
Dr J.B. Katz
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
2.5.4. Epizootic lymphangitis
Dr J. Picard
Department of Veterinary Tropical Diseases,
Faculty of Veterinary Science, University of
Pretoria, Private Bag X04, Onderstepoort 0110,
South Africa.
2.5.5. Equine encephalomyelitis
(Eastern and Western)
2.5.6. Equine infectious anaemia
Dr E.N. Ostlund
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
2.5.7. Equine influenza
Prof. Ann Cullinane
Irish Equine Centre, Johnstown, Naas,
Co. Kildare, Ireland.
2.5.8. Equine piroplasmosis
Dr T. de Waal
University College Dublin, School of Agriculture,
Food Science and Veterinary Medicine, Veterinary
Sciences Centre, Belfield, Dublin 4, Ireland.
2.5.9. Equine rhinopneumonitis
Dr G.P. Allen (deceased)
Department of Veterinary Science,
College of Agriculture, University of Kentucky, 108
M.H. Gluck Equine Research Center, Lexington,
Kentucky 40546-0099, USA.
2.5.10. Equine viral arteritis
Dr P.J. Timoney
University of Kentucky, Department of Veterinary
Science, 108 Gluck Equine Research Center,
Lexington, Kentucky 40546-0099, USA.
2.5.11. Glanders
Dr H. Neubauer
Friedrich-Loeffler Institut, Institut für Bakterielle
Infektionen und Zoonosen, Naumburger Strasse
96a, 07743 Jena, Germany.
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OIE Terrestrial Manual 2012
Contributors
2.5.12. Horse mange
See chapter 2.9.8.
2.5.13. Venezuelan equine encephalomyelitis
Dr E.N. Ostlund
USDA, APHIS, National Veterinary Services
Laboratories, P.O. Box 844, Ames, Iowa 50010,
USA.
2.6.1. Myxomatosis
Prof. S. Bertagnoli
École Nationale Vétérinaire, 23 Chemin de
Capelles, BP 87614, 31076 Toulouse Cedex 03,
France.
2.6.2. Rabbit haemorrhagic disease
Dr A. Lavazza & Dr L. Capucci
Istituto Zooprofilattico Sperimentale della
Lombardia e dell’Emilia Romagna, Via Bianchi
7/9, 25124 Brescia, Italy.
2.7.1. Border disease
Dr P.F. Nettleton & Dr K. Willoughby
Moredun Research Institute, International
Research Centre, Pentlands Science Park, Bush
Loan, Penicuik EH26 0PZ, Scotland, UK.
2.7.2. Caprine and ovine brucellosis (excluding
Brucella ovis)5
Dr B. Garin-Bastuji
EU Community/OIE & FAO Reference Laboratory
for Brucellosis, Unité Zoonoses Bactériennes,
Anses, 94706 Maisons-Alfort Cedex, France.
Dr J.M. Blasco
Centro de Investigación y Tecnología
Agroalimentaria de Aragón (CITAA), Apartado
727, 50080 Zaragoza, Spain.
2.7.3/4. Caprine arthritis/encephalitis & Maedi-visna
Dr D. Knowles & Dr L.M. Herrmann
USDA- ARS, Animal Disease Research Unit,
3003 ADBF, Washington State University,
Pullman, Washington 99164-6630, USA.
2.7.5. Contagious agalactia
Dr R. Nicholas
AHVLA Weybridge, New Haw, Addlestone,
Surrey KT15 3NB, UK.
Dr G.R. Loria
Istituto Zooprofilattico Sperimentale della Sicilia,
Palermo, Italy.
2.7.6. Contagious caprine pleuropneumonia
Dr F. Thiaucourt
CIRAD-EMVT, Campus international de
Baillarguet, Montferriez-sur-Lez, B.P. 5035, 34032
Montpellier Cedex 1, France.
2.7.7. Enzootic abortion of ewes (ovine chlamydiosis)
Dr K. Sachse
Friedrich-Loeffler-Institut, Federal Research
Institute for Animal Health, Institute of Molecular
Pathogenesis, Naumburger Str. 96a, 07743 Jena,
Germany.
Dr D. Longbottom
Moredun Research Institute, International
Research Centre, Pentlands Science Park
Bush Loan, Penicuik EH26 0PZ, UK.
2.7.8. Nairobi sheep disease
5
See chapter 2.9.1.
All four brucellosis chapters were updated by consensus of the OIE ad hoc Group on Brucellosis.
OIE Terrestrial Manual 2012
xxxi
Contributors
2.7.9. Ovine epididymitis (Brucella ovis)6
Dr B. Garin-Bastuji
EU Community/OIE & FAO Reference Laboratory
for Brucellosis, Unité Zoonoses Bactériennes,
Anses, 94706 Maisons-Alfort Cedex, France.
Dr J.M. Blasco
Centro de Investigación y Tecnología
Agroalimentaria de Aragón (CITAA), Apartado
727, 50080 Zaragoza, Spain.
2.7.10. Ovine pulmonary adenocarcinoma
(adenomatosis)
Dr M.J. Sharp
AHVLA, Lasswade Laboratory, Pentlands Science
Park, Bush Loan, Penicuik EH26 0PZ, Scotland,
UK.
2.7.11. Peste des petits ruminants7
Dr A. Diallo
FAO/IAEA Agriculture and Biotechnology
Laboratory, International Atomic Energy Agency,
A-2444 Seibersdorf, Austria.
2.7.12. Salmonellosis (S. abortusovis)
See chapter 2.9.9.
2.7.13. Scrapie
Dr M.M. Simmons, Mr M. Stack, Dr T. Konold &
Dr D. Matthews
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
2.7.14. Sheep pox and goat pox
Dr E. Tuppurainen
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Woking, Surrey GU24 0NF,
UK.
2.8.1. African swine fever
Dr C.A.L. Oura (formerly)
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Surrey GU24 0NF, UK.
Dr M. Arias
Centro de Investigación en Sanidad Animal
(CISA-INIA), Valdeolmos, 28130 Madrid, Spain.
6
7
8
2.8.2. Atrophic rhinitis of swine
Dr K.B. Register
USDA, ARS, National Animal Disease Center,
2300 Dayton Avenue, Ames, Iowa 50010, USA.
2.8.3. Classical swine fever (hog cholera)
Dr T. Drew
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
2.8.4. Nipah virus encephalitis
See chapter 2.9.6.
2.8.5. Porcine brucellosis8
Dr S. Olsen,
USDA, ARS, National Animal Disease Center,
2300 Dayton Avenue, Ames, Iowa 50010, USA.
2.8.6. Porcine cysticercosis
See chapter 2.9.5.
All four brucellosis chapters were updated by consensus of the OIE ad hoc Group on Brucellosis.
This chapter was updated by consensus of the OIE ad hoc Group on Peste des Petits Ruminants.
All four brucellosis chapters were updated by consensus of the OIE ad hoc Group on Brucellosis.
xxxii
OIE Terrestrial Manual 2012
Contributors
2.8.7. Porcine reproductive and respiratory syndrome
Dr L.R. Ludemann
USDA, APHIS, Center for Veterinary Biologics,
Laboratory, P.O. Box 844, Ames, Iowa 50010,
USA.
Dr K.M. Lager
Virus and Prion Diseases of Livestock Research
Unit, National Animal Disease Center, USDAARS, Ames, Iowa 50010, USA.
2.8.8. Swine influenza
Dr S.L. Swenson
National Veterinary Services Laboratories, P.O.
Box 844, Ames, Iowa 50010, USA.
2.8.9. Swine vesicular disease
Dr D. Paton
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Surrey GU24 0NF, UK.
Dr E. Brocchi,
Istituto Zooprofilattico Sperimentale della e
dell'Emilia Romagna (IZSLER), Via A. Bianchi
No. 9, 25124 Brescia, Italy.
Mr N. Knowles
2.8.10. Teschovirus encephalomyelitis (previously
Institute for Animal Health, Pirbright Laboratory,
enterovirus encephalomyelitis or Teschen/Talfan
Ash Road, Pirbright, Woking, Surrey GU24 0NF,
disease)
UK.
2.8.11. Transmissible gastroenteritis
Dr L.J. Saif
The Ohio State Universtiy, Ohio Agricultural
Research and Development Center, Food Animal
Health Research Program, 1680 Madison
Avenue, Wooster, Ohio 44691-4096, USA.
2.9.1. Bunyaviral diseases of animals (excluding
Rift Valley fever)
Dr G.H. Gerdes (retired)
Onderstepoort Veterinary Institute, Private Bag
X05, Onderstepoort 0110, South Africa.
2.9.2. Camelpox
Dr H. Elliott
International Animal Health Division , DEFRA,
1A Page Street, London SW1P 4PQ, UK.
Dr E. Tuppurainen
Institute for Animal Health, Pirbright Laboratory,
Ash Road, Pirbright, Surrey GU24 0NF, UK.
2.9.3. Campylobacter jejuni and C. coli
Prof. J.A. Wagenaar
Utrecht University, Faculty of Veterinary Medicine,
P.O. Box 80.165, 3508 TD Utrecht, The
Netherlands.
Dr W.F. Jacobs-Reitsma
RIKILT Institute of Food Safety, Wageningen-UR
P.O. Box 230, 6700 AE Wageningen, The
Netherlands.
2.9.4. Cryptosporidiosis
Prof. H. Smith
Scottish Parasite Diagnostic Laboratory, Stobhill
General Hospital, Glasgow G21 3UW, UK.
2.9.5. Cysticercosis
Dr S. Lloyd
Department of Clinical Veterinary Medicine,
University of Cambridge, Madingley Road,
Cambridge CB3 0ES, UK.
OIE Terrestrial Manual 2012
xxxiii
Contributors
2.9.6. Hendra and Nipah virus diseases
Dr P. Daniels
Australian Animal Health Laboratory, CSIRO
Livestock Industries, 5 Portarlington Road,
Geelong, Victoria 3220, Australia.
Dr M. Narasiman
Veterinary Research Institute, 59, Jalan Sultan
Azlan Shah, 31400 Ipoh, Perak, Malaysia.
Dr C.G. Gay
ARS, Research, Education, and Economics,
USDA, 5601 Sunnyside Avenue, Beltsville,
Maryland 20705-5148, USA.
Dr J.A. Roth
Center for Food Security and Public Health,
Institute for International Cooperation in Animal
Biologics
College of Veterinary Medicine, Iowa State
University, Ames, Iowa 50011, USA.
Dr H.M. Weingartl
Special Pathogens Unit, National Centre for
Foreign Animal Disease, Canadian Food
Inspection Agency, 1015 Arlington St., Winnipeg,
MB, R3E 3M4, Canada
2.9.7. Listeria monocytogenes
Dr J. Lopez (retired)
Canadian Food Inspection Agency,
National Centre for Foreign Animal Disease,
1015 Arlington Street, Winnipeg, Manitoba
R3E 3M4, Canada.
2.9.8. Mange
Dr J.L. Schlater & Dr J.W. Mertins
Parasitology and Clinical Pathology Section,
Pathobiology Laboratory, National Veterinary
Services Laboratories, USDA, APHIS, VS, P.O.
Box 844, Ames, Iowa 50010, USA.
2.9.9. Salmonellosis
Dr R. Davies
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
2.9.10. Toxoplasmosis
Dr D. Buxton & Dr S.W. Maley
Moredun Research Institute, Pentlands Science
Park, Bush Loan, by Edinburgh EH26 0PZ,
Scotland, UK.
2.9.11. Verocytotoxigenic Escherichia coli
Dr F.A. Clifton-Hadley
AHVLA Weybridge, New Haw, Addlestone, Surrey
KT15 3NB, UK.
2.9.12. Zoonoses transmissible from non-human
primates
FELASA Working Group on Non-Human Primate
Health: H. Weber (Convenor), E. Berge, J. Finch,
P. Heidt, F.-J. Kaup, G. Perretta, B. .Verschuere &
S. Wolfensohn
FELASA, BCM Box 2989, London WC1N 3XX,
UK.
Guideline 3.1. Laboratory methodologies for bacterial
antimicrobial susceptibility testing
Dr D. White
US Food and Drug Administration, Center for
Veterinary Medicine, Office of Research, 8401
Muirkirk Road, Laurel, MD 20708, USA.
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OIE Terrestrial Manual 2012
Contributors
Guideline 3.2. Biotechnology in the diagnosis of
infectious diseases
Dr G. Viljoen, Dr I. Naletoski & Dr A. Diallo
Joint FAO/IAEA Division of Nuclear Techniques in
Food and Agriculture, Department of Nuclear
Sciences and Applications, International Atomic
Energy Agency, A-2444 Seibersdorf, Austria.
Guideline 3.3. The Application of Biotechnology to the
Development of Veterinary Vaccines
Dr A.A. Potter, Dr V. Gerdts, Dr G. Mutwiri,
Dr S. Tikoo & De S. van Drunen Littelvan den Hurk
Vaccine and Infectious Disease Organization,
120 Veterinary Road, Saskatoon S7N 5E3,
Canada.
Guideline 3.4. The role of official bodies in the
international regulation of veterinary biologicals
Dr Ph. Vannier (retired)
Anses Ploufragan, Laboratoire d’études et de
recherches avicoles et porcines, Zoopôle des
Côtes d’Armor-Les Croix, BP 53, 22440
Ploufragan, France.
Dr R. Hill
Center for Veterinary Biologics, USDA, APHIS,
Veterinary Services, P.O. Box 844, Ames
Iowa 50010, USA.
Dr O. Itoh
National Veterinary Assay Laboratory, JMAFF,
1-15-1 Tokura, Kokubunji, Tokyo 185-8511,
Japan.
Dr C. Lambert
Anses Fougères, Agence nationale du
médicament vétérinaire, B.P. 203, 35302
Fougères Cedex, France.
OIE Terrestrial Manual 2012
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2012
SECTION 2.5.
EQUIDAE
CHAPTER 2.5.1.
AFRICAN HORSE SICKNESS
SUMMARY
Description of the disease: African horse sickness (AHS) is an infectious but noncontagious viral
disease affecting all species of equidae caused by an Orbivirus of the family Reoviridae and
characterised by alterations in the respiratory and circulatory functions. AHS is transmitted by at
least two species of Culicoides. Nine different serotypes have been described.
All serotypes of AHS occur in eastern and southern Africa. AHS serotypes 9, 4 and 2 have been
found in North and West Africa from where they occasionally spread into countries surrounding the
Mediterranean. Examples of outbreaks that have occurred outside Africa are: in the Middle East
(1959–1963), in Spain (serotype 9, 1966, serotype 4, 1987–1990), and in Portugal (serotype 4,
1989).
Laboratory diagnosis of AHS is essential. Although the clinical signs and lesions are characteristic,
they can be confused with those of other equine diseases.
As a viral disease, the laboratory diagnosis of AHS can be based on the identification of infectious
virus, virus nucleic acid, viral antigens or specific antibodies. Over the past few years, a wide
variety of laboratory tests have been adapted for the detection of both AHS virus (AHSV) and
specific antibodies.
Identification of the agent: it is particularly important to perform virus isolation and serotyping
whenever outbreaks occur outside the enzootic regions in order to choose a homologous serotype
for the vaccine.
AHSV can be isolated from blood collected during the early febrile stage. For virus isolation, the
other tissues of choice for diagnosis are spleen, lung, and lymph nodes, collected at necropsy.
Sample preparations can be inoculated in cell cultures, such as baby hamster kidney-21 (BHK-21),
monkey stable (MS), African green monkey kidney (Vero) or insect cells (KC), intravenously in
embryonated eggs, and intracerebrally in newborn mice. Several enzyme-linked immunosorbent
assays (ELISAs) for the rapid detection of AHSV antigen in blood, spleen tissues and supernatant
from infected cells have been developed. Identification of AHSV RNA has also been achieved using
a reverse-transcription polymerase chain reaction method. Virus isolates can be serotyped by a
type-specific serological test such as virus neutralisation (VN) and by reverse-transcription
polymerase chain reaction and sequencing.
Serological tests: Horses that survive natural infection develop antibodies against the infecting
serotype of AHSV within 8–12 days post-infection. This may be demonstrated by several
serological methods, such as complement fixation test, ELISA, immunoblotting and VN. The latter
test is used for serotyping.
Requirements for vaccines: Attenuated (monovalent and polyvalent) live vaccines for use in
horses, mules and donkeys, are currently commercially available. Subunit vaccines have been
evaluated experimentally.
OIE Terrestrial Manual 2012
819
Chapter 2.5.1. — African horse sickness
A. INTRODUCTION
African horse sickness (AHS) (Peste equina africana, Peste equine) is an infectious, non-contagious arthopodborne disease of equidae, caused by a double-stranded RNA Orbivirus belonging to the family Reoviridae. The
genus Orbivirus also includes bluetongue virus and epizootic haemorrhagic disease virus, which have similar
morphological and biochemical properties with distinctive pathological and antigenic properties as well as host
ranges. Nine antigenically distinct serotypes of AHSV have been identified by virus neutralisation; some crossreaction has been observed between 1 and 2, 3 and 7, 5 and 8, and 6 and 9, but no cross-reactions with other
known orbiviruses occur.
The virion is an unenveloped particle of a size around 70 nm. The genome of AHS virus (AHSV) is composed of
ten double-stranded RNA segments, which encode seven structural proteins (VP1-7), most of which have been
completely sequenced for AHSV serotypes 4, 6 and 9 (Roy et al., 1991; Venter et al., 2000; Williams et al., 1998)
and four nonstructural proteins (NS1, NS2, NS3, NS3A) (Grubman & Lewis, 1992; Laviada et al., 1993). Proteins
VP2 and VP5 form the outer capsid of the virion, and proteins VP3 and VP7 are the major inner capsid proteins.
Proteins VP1, VP4 and VP6 constitute minor inner capsid proteins. The NS3 proteins are the second most
variable AHSV proteins (Van Niekert et al., 2001), the most variable being the major outer capsid protein, VP2.
This protein, VP2, is the determinant of AHSV serotypes and, together with VP5, the target for virus neutralisation
activity (Martinez-Torrecuadrada et al., 2001). At least two field vectors are involved in the transmission of the
virus: Culicoides imicola and C. bolitinos.
AHS is enzootic in sub-Saharan Africa, although occasional outbreaks have occurred in northern Africa (1965,
1989–1990), the Middle East (1959–1961), and in Europe (Spain, 1966, 1987–1990 and Portugal, 1989)
(Sanchez-Vizcaíno, 2004).
There are four classical clinical forms of AHS: pulmonary, cardiac, mixed, and horse sickness fever. The
peracute, pulmonary form occurs in fully susceptible animals and has a short course, often only a few hours, and
a high mortality rate. The animal exhibits respiratory distress, an extended head and neck, and profuse sweating.
Terminally, froth exudes from the nostrils. The cardiac, oedematous form has a more subacute course with
mortality reaching 50%. The head and neck may show severe swelling that can extend down to the chest.
Swelling of the supraorbital fossae is characteristic and may include conjunctival swelling with petechiae.
Paralysis of the oesophagus may result in aspiration pneumonia and sublingual haemorrhages are always a poor
prognostic sign. The mixed, acute form is most commonly seen and has feature of both the cardiac and
pulmonary forms. Mortality can reach 70%. Horsesickness fever is an often overlooked, mild form of the disease
and is seen in resistant equidae such as zebra and donkeys (Coetzer & Guthrie, 2005).
The disease has both a seasonal (late summer/autumn) and a cyclical incidence with major epizootics in southern
Africa during warm-phase events (Baylis et al., 1999). Mortality due to AHS is related to the species of equidae
affected and to the strain or serotype of the virus. Among equidae, horses are the most susceptible to AHS with a
mortality rate of 50–95%, followed by mules with mortality around 50%. In enzootic regions of Africa, donkeys are
very resistant to AHS and experience only subclinical infections. In European and Asian countries, however,
donkeys are moderately susceptible and have a mortality rate of 10%. Zebras are also markedly resistant with no
clinical signs, except fever, and may have extended viraemia (up to 40 days).
A laboratory diagnosis is essential to establish a correct and confirmatory diagnosis. Although some clinical signs
and lesions are characteristic, AHS can be confused with other diseases. For example, the supraorbital swelling,
which is often present in horses with subacute AHS, is, in combination with an appropriate history, sufficient for a
tentative diagnosis. Other signs and lesions are less specific for AHS, and other diseases such as equine
encephalosis, equine infectious anaemia, equine morbillivirus pneumonia, equine viral arteritis, babesiosis and
purpura haemorrhagica should be excluded (OIE Technical Disease Cards: http://www.oie.int/en/animal-health-inthe-world/technical-disease-cards/).
Attenuated (monovalent and polyvalent) live vaccines for use in horses, mules and donkeys, are currently
commercially available.
There is no evidence that humans can become infected with any field strain of AHSV, either through contact with
naturally or experimentally infected animals or by virus manipulation in laboratories.
B. DIAGNOSTIC TECHNIQUES
Several techniques are already available for AHS viral identification ranging from the rapid capture (indirect
sandwich) enzyme-linked immunosorbent assay (ELISA), using either polyclonal antibodies (PAbs) or monoclonal
antibodies (MAbs), to the polymerase chain reaction (PCR) test, including a new reverse-transcription (RT) PCR
for discrimination of the nine AHSV serotypes or cell culture and inoculation of newborn mice. If possible more
than one test should be performed to diagnose an outbreak of AHS, especially the index case. The initial test can
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Chapter 2.5.1. — African horse sickness
be a quick test such as ELlSA or PCR, followed by virus isolation in tissue culture. Virus neutralisation (VN) for
serotype identification or RT-PCR with sequencing should be performed as early in the outbreak as possible so
that the serotype can be identified and the correct vaccine selected.
At present, there are no international standards for viruses or diagnostic reagents, and there is no standard
methodology for the identification of AHSV. However, a viral and antibody panel has been evaluated, and
comparative studies between different ELISAs for AHSV antigen and antibody determination have been carried
out in different laboratories, including in the EU Reference Laboratory for AHS. The results have demonstrated a
high level of correlation for both antigen and antibody determination with an in-house test and commercial kits
(Rubio et al., 1998; Villalba, 2009). Similar studies have been conducted with several RT-PCR assays (Aquero,
2009) also providing a high level of correlation.
A very important aspect of the diagnosis is the selection of samples and their safe transportation to the laboratory.
1.
Identification of the agent
a)
Virus isolation
Unclotted whole blood collected during the early febrile stage of the disease from sick animals, as well as
small pieces (2–4 g) of spleen, lung and lymph nodes from animals that have died, are the samples of
choice for diagnosis. Samples should be kept at 4°C during transportation and short-term storage prior to
processing.
•
Cell culture
Successful direct isolation of AHSV has been performed on baby hamster kidney (BHK-21), monkey stable
(MS) and African green monkey kidney (Vero) mammalian cell lines and on Culicoides and mosquito insect
cell lines. Blood samples collected in an appropriate anticoagulant can be used undiluted as the inoculum.
After 15–60 minutes of adsorption at ambient temperature or at 37°C, the cell cultures are washed and
maintenance medium is added. Alternatively and more commonly, the blood is washed, lysed and diluted
1/10. This procedure removes unwanted antibody, which could neutralise free virus, and promotes release
of virus associated with the red blood cell membranes. When tissue samples, such as spleen, lung, etc., are
used, a 10% tissue suspension is prepared in phosphate buffered saline (PBS) or cell culture medium,
containing antibiotics.
A cytopathic effect (CPE) may appear between 2 and 10 days post-infection with mammalian cells. Three
blind passages should be performed before considering the samples to be negative. No CPE is observed in
insect cells but the presence of the virus can be detected in the supernatant after 5–7 days by real-time RTPCR. Supernatant from infected insect cells can then be passed onto mammalian cells, which will show CPE
after one or two passages.
•
Newborn mice
This method of isolation of AHSV involves the intracerebral inoculation of two families of 1–3-day-old mice.
In positive cases, animals develop nervous signs between 3 and 15 days post-inoculation. The brains from
sick animals may be collected, homogenised and re-inoculated intracerebrally into at least six 1–3-day-old
mice. This second passage should present a shortened incubation period (2–5 days) and 100% mortality.
Virus may be typed directly from mouse brain by conventional neutralisation (VN) or by RNA extraction and
sequencing.
b)
Nucleic acid methods
•
Reverse-transcription polymerase chain reaction (an alternative test for international trade)
The reverse-transcription polymerase chain reaction (RT-PCR) is a highly sensitive technique that allows the
detection of a very low number of copies of RNA molecules, but this does not necessarily indicate the
presence of infectious virus. The RNA strand is first reverse transcribed into its complementary DNA
(cDNA), followed by amplification of the resulting DNA using polymerase chain reaction. This technique has
greatly improved the detection of AHSV-RNA by improving the sensitivity of detection and shortening the
time required for the diagnosis.
Several agarose gel-based RT-PCR assays for the specific detection of AHSV RNA have been described
targeted at viral segments 3, 7 or 8 (Aradaib, 2009; Bremer et al., 1998; Laviada et al., 1997; Sakamoto et
al., 1994; Stone-Marschat et al., 1994; Zientara et al., 1994). The most widely used method employs primers
corresponding to the 5’ end (nucleotides 1–21) and 3’ end (nucleotides 1160–1179) of RNA segment 7
amplifying the complete viral segment.
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Chapter 2.5.1. — African horse sickness
Real-time RT-PCR (rRT-PCR) methods for the highly sensitive and specific detection of AHSV RNA have
been recently developed based on the use of a pair of primers and a Taqman probe from conserved
sequences of viral segments 5 or 7 (Agüero et al.. 2008; Fernández-Pinero et al., 2009; Rodriguez-Sanchez
et al., 2008). A duplex RRT-PCR has also been described that targets segments 8 and 9 of the genome and
uses Taqman® probes (Quan et al., 2010). Although both gel-based and rRT-PCR procedures can detect
reference strains from the nine virus serotypes, rRT-PCR provides advantages over agarose gel-based RTPCR methods, with its faster analysis time, higher sensitivity, and suitability for high-throughput automation.
Nevertheless gel-based RT-PCR methods, particularly those amplifying long RNA fragments (Laviada et al.,
1997; Zientara et al., 1994), can be very useful in the further genetic characterisation of the virus by
sequencing of the amplicons.
Details of AHSV agarose gel-based RT-PCR and real-time RT-PCR are given below.
•
Extraction of viral RNA
To assure a good reaction it is necessary to extract from the sample an AHSV RNA of high quality. The
extraction of nucleic acids from clinical samples can be performed by a variety of in-house and commercially
available methods.
An example of an in-house RNA extraction is given below:
i)
1 g of tissue sample is homogenised in 1 ml of denaturing solution (4 M guanidium thiocyanate, 25 mM
sodium citrate, 0.1 M 2-mercaptoethanol, 0.5% sarcosyl).
ii)
After centrifugation, 1 µg of yeast RNA, 0.1 ml of 2 M sodium acetate pH 4, 1 ml of phenol and 0.2 ml
of chloroform/isoamyl alcohol mixture (49/1) are added to the supernatant.
iii)
The suspension is vigorously shaken and cooled on ice for 15 minutes.
iv)
After centrifugation, the RNA present in the aqueous phase is phenol extracted, ethanol precipitated
and resuspended in sterile water
Commercial kits use different approaches for RNA isolation. Most are based on one of the following
procedures:
•
•
Phenol–chloroform extraction of nucleic acids;
•
Adsorption of nucleic acids to filter system;
•
Adsorption of nucleic acids to magnetic beads system.
Agarose gel-based RT-PCR procedure (Laviada et al., 1997; Zientara et al., 1994)
Denaturation of extracted RNA has to be performed prior to the RT-PCR procedure. The sequences of the
PCR primers used are 5’-GTT-AAA-ATT-CGG-TTA-GGA-TG-3’, which corresponds to the messenger RNA
polarity, and 5’-GTA-AGT-GTA-TTC-GGT-ATT-G-3’, which is complementary to the messenger RNA
polarity. All the components required for the reverse transcription and PCR are included in the reaction tube
containing the denatured RNA. A one-step RT-PCR is carried out by incubating in a thermocycler as follows:
45 minutes to 1 hour at 37–55°C, 5–10 minutes at 95°C, then 40 cycles of: 94–95°C for 1 minute, 55°C for
1–1.5 minutes, 70–72°C for 2–2.5 minutes, followed by a final extension step of 7–8 minutes at 70–72°C.
Analysis of the PCR products is carried out by agarose gel electrophoresis. AHS-positive samples will
resolve in a 1179 base-pair band that can be used as template in the sequencing reaction, using
independently the PCR primers, for obtaining the nucleotide sequence of viral segment 7.
•
Real-time RT-PCR Procedure (Agüero et al., 2008)
This group-specific real-time RT-PCR has been employed with good results by the participating national
reference laboratories of the European Union (EU) Member States in a proficiency test organised by the EU
reference laboratory for AHS (Agüero, 2009). The rRT-PCR is carried out as follows:
822
i)
2 µl of isolated RNA is mixed with forward (5’-CCA-GTA-GGC-CAG-ATC-AAC-AG-3’) and reverse (5’CTA-ATG-AAA-GCG-GTG-ACC-GT-3’) primers (2.5 µl of each primer at 8 µM) and RNAse-free water
up to 7 µl.
ii)
This mixture is denatured by heating at 95°C for 5 minutes, followed by rapid cooling on ice.
iii)
cDNA synthesis and hot-start PCR amplification are carried out in one-step, in a volume of 20 µl
containing the denatured RNA-primers mixture, 0.1 µl of the fluorogenic MGB-TaqMan probe (5’-FAMGCT-AGC-AGC-CTA-CCA-CTA-MGB-3’, probe was labelled in 5’ with 6-carboxyfluorescein, FAM, and
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Chapter 2.5.1. — African horse sickness
in 3’ with a non-fluorescent quencher bound to an MGB group) at 50 µM (final concentration: 0.25 µM),
adequate buffer and enzymes (RTase and DNA polymerase at a concentration recommended by the
manufacturer).
iv)
Amplification conditions consist of a first reverse-transcription step at 48°C for 25 minutes, followed by
10 minutes at 95°C (‘hot start’) and 40 cycles of 15 seconds at 95°C, 35 seconds at 55°C and
30 seconds at 72°C (or 40 cycles at 97°C for 2 seconds and 55°C for 30 seconds if reagents and
thermocycler allowing fast reactions are used). Fluorescence data are acquired at the end of the 55°C
step.
v)
Samples are considered positives if the fluorescence increases significantly over the base level. If no
fluorescence is detected during the whole real-time RT-PCR, samples are considered negative.
Inactivated virus of serotypes 1-9 reference strains can be obtained from the OIE Reference Laboratory in
Spain in order to set up the RT-PCR detection method.
c)
AHSV typing
Until recently, the VN test has been the method of choice for typing as well as the ‘gold’ standard test for
identifying AHSV isolated from the field using type-specific antisera (Verwoerd, 1979). This technique takes
5 or more days before results are obtained. The development of type-specific gel-based RT-PCR (Sailleau
et al., 2000), and real-time RT-PCR using hybridisation probes (Koekemoer, 2008) for identification and
differentiation of AHSV genotypes, provides a rapid typing method for AHSV in tissue samples and blood.
There is a good correlation between the results obtained with the type-specific RT-PCR and the VN test,
however, the sensitivity of these assays is lower than that obtained with the diagnostic group-specific realtime RT-PCR (Agüero et al., 2008). Type-specific rRT-PCR assays based on the use of Taqman-MGB
probes have been developed more recently (Tena, 2009) and have a similar sensitivity to the group-specific
rRT-PCR. However, the genetic variation that may appear over time in the AHSV genome, in particular in
the VP2 coding region, where specific primers/probes for typing assays have to be designed, makes the
detection of all genetic variants within each serotype by this type of technique difficult (Koekemoer, 2008).
Therefore, although molecular methods are able to rapidly type AHSV in many positive field samples, VN
should be kept as the gold standard for serotyping AHSV isolates.
Typing of the nine AHSV types has also been carried out with probes developed from a set of cloned fulllength VP2 genes (Koekemoer et al., 2000). This technique can be used as an alternative to PCR
amplification of genome segment 2.
2.
Serological tests
Indirect and competitive blocking ELISAs using either soluble AHSV antigen or a recombinant protein VP7
(Hamblin et al., 1990; Laviada et al., 1992b; Maree & Paweska, 2005) have proved to be good methods for the
detection of anti-AHSV group-reactive antibodies, especially for large-scale investigations (Rubio et al., 1998).
Both of these tests have been recognised by the European Commission (2002). The competitive blocking ELISA
can also be used for testing wildlife as species-specific anti-globulin is not required with this method. An
immunoblotting test has also been adapted for anti-AHS antibody determination (Laviada et al., 1992b), which is
especially suitable for small numbers of sera. The complement fixation (CF) test has been widely used, but some
sera are anti-complementary, particularly donkey and zebra sera.
a)
Blocking enzyme-linked immunosorbent assay (a prescribed test for international trade)
The competitive blocking ELISA technique aims at detecting specific antibodies against AHSV, present in
any equine species. VP7 is the main antigenic protein within the molecular structure of AHSV and it is
conserved across the nine AHSV serotypes. An MAb directed against VP7 is used in this test, allowing high
sensitivity and specificity. Moreover, other species apart from horses (e.g. donkeys, zebra, etc.) can be
tested thus preventing the problem of specificity experienced occasionally using the indirect ELISAs. VP7
recombinant antigen is non-reactive, which provides a high level of security (European Commission, 2002).
The principle of this test is to block the specific reaction between the recombinant VP7 protein absorbed on
an ELISA plate and a conjugated MAb against VP7. AHSV antibodies present in a suspect serum sample
will block this reaction. A decrease in the amount of colour is evidence of the presence of AHSV antibodies
in the serum sample.
•
Test procedure
i)
Solid phase: coat ELISA plates (e.g. high adsorption capacity Nunc Maxisorb) with 50–100 ng of
recombinant AHSV-4 VP7 diluted in carbonate/bicarbonate buffer, pH 9.6. Incubate overnight at 4°C.
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Chapter 2.5.1. — African horse sickness
ii)
Wash the plates three times with PBS 0.1× containing 0.135 M NaCl and 0.05% (v/v) Tween 20
(washing solution). Gently tap the plates on to absorbent material to remove any residual wash.
iii)
Test samples: serum samples to be tested, and positive and negative control sera (if not ready to use
by kit manufacturer), are diluted 1/5 in diluent containing 0.35 M NaCl, 0.05% Tween 20; and
0.1% Kathon, 100 µl per well. Incubate for 1 hour at 37°C.
iv)
Wash the plates five times with PBS 0.1× containing 0.135 M NaCl and 0.05% (v/v) Tween 20
(washing solution). Gently tap the plates on to absorbent material to remove any residual wash.
v)
Conjugate: dispense 100 µl/well of horseradish peroxidise-conjugated MAb anti-VP7. This MAb should
be previously diluted 1/5000–1/15000 in a 1/1 solution of StabiliZyme Select® Stabilizer (SurModics.
Reference: SZ03) in distilled water. Incubate for 30 minutes at 37°C.
vi)
Wash the plates as described in step iv.
vii)
Substrate/chromogen: add 100 µl/well of 1/10 diluted ABTS substrate solution, 5 mg/ml substrate
buffer (0.1 M phosphate/citrate buffer, pH 4, containing 0.03% H2O2) and incubate for 10 minutes at
room temperature. Colour development is stopped by adding 100 µl/well of 2% (w/v) of SDS.
viii) Read the plates at 405 nm.
ix)
Interpretation of results: determine the blocking percentage (BP) of each sample by applying the
following formula:
BP=
Abs(control–) – Abs(sample)
Abs(control–) – Abs(control+)
× 100
Samples showing BP value lower than 45% are considered negative for antibodies to AHSV. Samples
showing BP value higher than 50% are considered positive for antibodies to AHSV. Samples with BP value
between 45% and 50% are considered doubtful and must be retested. If the result is the same, resample
and test 2 weeks later.
b)
Indirect enzyme-linked immunosorbent assay (a prescribed test for international trade)
The recombinant VP7 protein has been used as antigen for AHSV antibody determination with a high degree
of sensitivity and specificity (Laviada et al., 1992b; Wade-Evans et al., 1993). Other advantages of this
antigen are its stability and its lack of infectivity. The conjugate used in this method is a horseradish
peroxidase anti-horse gamma-globulin reacting with horse, mules and donkeys. The method described by
Maree & Paweska (2005) uses protein G as conjugate that also reacts with zebra serum.

Test procedure
i)
Solid phase: Coat ELISA plates (e.g. high adsorption capacity Nunc Maxisorb) with recombinant
AHSV-4 VP7 diluted in carbonate/bicarbonate buffer, pH 9.6. Incubate overnight at 4°C.
ii)
Wash the plates five times with distilled water containing 0.01% (v/v) Tween 20 (washing solution).
Gently tap the plates on to absorbent material to remove any residual wash.
iii)
Block the plates with PBS, pH 7.2 + 5% (w/v) skimmed milk, 200 µl/well, for 1 hour at 37°C.
iv)
Remove the blocking solution and gently tap the plates on to absorbent material.
v)
Test samples: Serum samples to be tested, and positive and negative control sera, are diluted 1/25 in
PBS + 5% (w/v) skimmed milk + 0.05% (v/v) Tween 20, 100 µl per well. Incubate for 1 hour at 37°C.
For titration, add twofold dilution series from 1/25 (100 µl/well), one serum per plate column, and do the
same with positive and negative controls. Incubate for 1 hour at 37°C.
vi)
Wash the plates as described in step ii.
vii)
Conjugate: Dispense 100 µl/well of horseradish peroxidase conjugated anti-horse gamma-globulin
diluted in PBS + 5% milk + 0.05% Tween 20, pH 7.2. Incubate for 1 hour at 37°C or protein A
peroxidase (Maree & Paweska, 2005).
viii) Wash the plates as described in step ii.
ix)
824
Substrate: Add 200 µl/well of substrate solution (10 ml DMAB + 10 ml of MBTH + 5 µl H2O2). Colour
development is stopped by adding 50 µl of 3 N H2SO4 after approximately 5–10 minutes (before the
negative control begins to be coloured). Other substrates such as ABTS (2,2’-azino-di-[3-ethylbenzothiazoline]-6-sulphonic acid), TMB (tetramethyl benzidine), or OPD (orthophenyldiamine) can
also be used.
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Chapter 2.5.1. — African horse sickness
c)
x)
Read the plates at 600 nm (or 620 nm).
xi)
Interpretation of results: Calculate the cut-off value by adding 0.6 to the value of the negative control.
(0.06 is the standard deviation derived with a group of 30 negative sera) Test samples giving
absorbance values lower than the cut-off are regarded as negative. Test samples giving absorbance
values greater than the cut-off + 0.15 are regarded as positive. Test samples giving intermediate
absorbance values are doubtful and a second technique must be employed to confirm the result.
Complement fixation (a prescribed test for international trade)
The CF test has been used extensively in the past, but currently its use is decreasing and has been replaced
in many laboratories by ELISA as a screening technique. This progressive replacement is because of the
higher sensitivity and degree of standardisation of ELISA as well as a significant number of sera with anticomplementary activity. Nevertheless the CF test is a useful tool in endemic areas for the demonstration and
titration of group-specific IgM antibodies against AHSV notably following a recent infection or vaccination.

Reagents
i)
Veronal buffered saline containing 1% gelatin (VBSG).
ii)
Serum samples, free from erythrocytes, must be heat inactivated: horse serum at 56°C, zebra serum at
60°C and donkey serum at 62°C, for 30 minutes.
iii)
The antigen is a sucrose/acetone extract of AHSV-infected mouse brain. The control antigen is
uninfected mouse brain, extracted in the same way. In the absence of an international standard serum,
the antigen should be titrated against a locally prepared positive control serum. In the test, four to eight
units are used. The antigen may also be obtained by inoculation of the virus in suitable cell culture (see
Section B.1 above).
iv)
The complement is a normal guinea-pig serum.
v)
The haemolysin is a hyperimmne rabbit serum against sheep red blood cells (SRBCs).
vi)
The SRBCs are obtained by aseptic puncture of the jugular vein and preserved in Alsever’s solution1 or
sodium citrate.
vii)
The haemolytic system (HS) is prepared by diluting the haemolysin to contain two haemolytic doses
and using this to sensitise washed SRBCs. The SRBCs are standardised to a 3% concentration.
viii) Control sera: A positive control serum is obtained locally and validated. Serum from a healthy antibodynegative horse is used as the negative control serum.

Test procedure
i)
The reaction is performed in 96-well round-bottom microtitre plates in a final volume of 100 µl/well or in
tubes if the macro-technique is used, at 4°C for 18 hours.
ii)
All the sera, samples and controls are diluted 1/5 in VBSG and 25 µl of each serum is added in
duplicate. A twofold dilution series of each serum is done from 1/5 to 1/180.
iii)
Add 25 µl of the antigen diluted according to the previous titration.
iv)
Add 25 µl of the complement diluted according to a previous titration.
v)
Incubate at 4°C for 18 hours.
vi)
25 µl of HS is added to all wells on the microtitre plate.
vii)
The plate is incubated for 30 minutes at 37°C.
viii) Plates are then centrifuged at 200 g, and the wells are scored for the presence of haemolysis. Control
of sera, complement, antigen and HS are used
1
ix)
Results are read using 50% haemolysis as the end point. The inverse of the highest dilution of serum
specifically fixing complement with the CF antigen is called the titre.
x)
A titre of 1/10 or more is positive, under 1/10 is negative.
20.5 g dextrose (114 mM), 7.9 g sodium citrate 2H2O (27 mM), 4.2 g NaCl (71 mM), H2O to 1 litre. Adjust to pH with 1 M
citric acid.
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Chapter 2.5.1. — African horse sickness
d)
Virus neutralisation (VN)
Serotype-specific antibody can be detected using the VN test (House et al., 1990). The VN test may have
additional value in epidemiological surveillance and transmission studies, mainly in endemic areas where
multiple serotypes are likely to be present.

VN Test procedure
i)
Stock virus is diluted to yield 100 TCID50 (50% tissue culture infective dose), with a range of 30–
300 TCID50, per 25 µl, and 25 µl is added to each of four microtitre wells containing 25 µl serum
dilutions. For screening, a final serum dilution of 1/10 is used. Doubling dilutions are used for titrations.
ii)
Serum/virus mixtures are incubated for 60 minutes at 37°C 5% CO2 and 95% humidity prior to the
addition of 0.1 ml of Vero cell suspension (200,000 cells/ml) to each test well.
iii)
A back titration of virus stock is prepared for each test using four wells per tenfold dilution, 25 µl per
well. Test plates are incubated at 37°C, 5% CO2, 95% humidity for 4–5 days, until the back titration
indicates that the stock virus contains 30–300 TCID50.
iv)
The plates are then fixed and stained in a solution of 0.15% (w/v) crystal violet in 2% (v/v)
glutaraldehyde and rinsed. Alternatively, they may be fixed with 70% ethanol and stained with 1% basic
fuschsin.
v)
The 50% end-point titre of the serum is calculated by the Spearman–Kärber method and expressed as
the negative log10.
C. REQUIREMENTS FOR VACCINES
1.
Background
a)
Rationale and intended use of the product
Polyvalent or monovalent live attenuated AHS vaccines, based on the selection in Vero cell culture of
genetically stable macroplaques, have been used for the control of AHSV in and out of Africa (Erasmus,
1976; Sanchez-Vizcaíno, 2004). Polyvalent vaccines are commercially available.
An inactivated monovalent (serotype 4) AHSV vaccine based on virus purification and inactivation with
formalin was produced commercially in the early 1990s (House et al., 1992), but is not available at the
present time. Subunit AHSV vaccines based on serotype 4 outer capsid protein VP2 and VP5 plus inner
capsid protein VP7, derived from single and dual recombinant baculovirus expression vectors have been
used experimentally in different combinations to immunise horses (Martinez et al., 1996). The protective
efficacy of VP2 in a subunit vaccine was also evaluated (Scanlen et al., 2002). However, these vaccines are
not commercially available.
2.
Outline of production and minimum requirements for conventional vaccines
At present only the live attenuated AHS vaccines (polyvalent or monovalent) are commercially available.
Guidelines for the production of veterinary vaccines are given in Chapter 1.1.6 Principles of veterinary vaccine
production. The guidelines given here and in chapter 1.1.6 are intended to be general in nature and may be
supplemented by national and regional requirements.
•
Live attenuated African horse sickness vaccine
a)
Characteristics of the seed
i)
Biological characteristics
The seed virus is prepared by selection in Vero cells of genetically stable large plaques from low
passage levels of AHSV. The plaque mutants are then further multiplied by three passages in Vero
cells. A large quantity of this antigen is lyophilised and stored at –20°C as seed stock antigen.
ii)
Quality criteria
The seed virus must be shown to be free of contaminating viruses, bacteria and mycoplasmas by the
appropriate techniques. The serotype identity of the seed virus is confirmed.
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Chapter 2.5.1. — African horse sickness
b)
Method of manufacture
i)
Procedure
At the onset of a production run, working antigens are produced from the seed stock antigen in either
BHK-21 or Vero cell cultures. The working antigens are tested for sterility, purity and identity and
should contain at least 1 × 106 plaque-forming units (PFU)/ml of infectious virus.
ii)
Requirements for substrates and media
Roller bottle cultures of Vero or BHK-21 cells are grown using gamma-irradiated bovine serum in the
growth medium. Once the cultures are confluent, the medium is poured off and the cells are seeded
with the working antigens. After 1 hour, maintenance medium is added to the cultures. Incubation is
continued at 37°C for 2–3 days. When the CPE is advanced, both cells and supernatant medium are
harvested. The products from the same serotype are pooled and stored at 4°C.
iii)
In-process control
The pooled harvests of the individual serotypes are tested for sterility and assayed for infectivity by
plaque titration on Vero cell cultures. The minimum acceptable titre is 1 × 106 PFU/ml.
Finally, two quadrivalent vaccines are constituted by mixing equal volumes of serotypes 1, 3, 4, 5 and
2, 6, 7, 8 respectively. Subsequently, AHSV serotype 5 was withdrawn from this vaccine. A monovalent
type can also be prepared. After addition of suitable stabiliser, the vaccine is distributed in 1.0 ml
volumes into glass vials and freeze-dried.
iv)
Final product batch test
Sterility
Following lyophilisation, five bottles of vaccine are selected at random and tested for sterility by
internationally accepted methods. Tests for sterility and freedom from contamination of biological
products are given in chapter 1.1.7.
Safety
Innocuity of a vaccine is determined by the inoculation of reconstituted vaccine into mice (0.25 ml
intraperitoneally), guinea-pig (1.0 ml intraperitoneally), and a horse (5.0 ml subcutaneously). All the
animals are observed daily for 14 days. The rectal temperature of the horse is taken twice daily for
14 days and should never exceed 39°C.
Batch potency
Potency is largely based on virus concentration in the vaccine.
The minimum immunising dose for each serotype is about 1 × 103 PFU/dose. The infectivity titre of the
final product is assayed by plaque titration in Vero cell cultures and should contain at least 1 ×
105 PFU/dose. The horse used for safety testing is also used for determining the immunogenicity of a
vaccine.
Serum samples are collected on the day of vaccination and 21 days later, and are tested for
neutralising antibodies against each serotype by the plaque-reduction test using twofold serum
dilutions and about 100 PFU of virus. The horse should develop a neutralising antibody titre of at least
20 against at least three of the four serotypes in the quadrivalent vaccine.
c)
Requirements for authorisation
No specific guideline is described for AHS vaccine. However a guideline is described in the EU for
Bluetongue virus under exceptional circumstances that could probably be used for AHS virus. This guideline
includes the minimum date requirements for the authorisation under exceptional circumstances for vaccine
production for emergency use against bluetongue virus (Regulation EC Nº726/2004, in particular Articles 38,
39 and 43 thereof and Article 26 of Direction 2001/82/EC), including guidance measures to facilitate the
rapid inclusion of new or different virus serotypes.
3.
Vaccines based on biotechnology
a)
Vaccines available and their advantages
None is available commercially. Experimental subunit vaccines have been described (Section C.1.a above).
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Chapter 2.5.1. — African horse sickness
b)
Special requirements for biotechnological vaccines, if any
None.
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National African Horse Sickness (AHS) Reference Laboratories. Brussels, Belgium 1 December 2009.
AGÜERO M., GÓMEZ-TEJEDOR C., ANGELES CUBILLO M., RUBIO C., ROMERO E. & JIMÉNEZ-CLAVERO A. (2008). Realtime fluorogenic reverse transcription polymerase chain reaction assay for detection of African horse sickness
virus. J. Vet. Diagn. Invest., 20, 325–328.
ARADAIB I.E. (2009). PCR detection of African horse sickness virus serogroup based on genome segment three
sequence analysis. J. Virol. Methods, 159 (1),1–5.
BAYLIS M., MELLOR P.S. & MEISWINKEL R. (1999). Horse sickness and ENSO in South Africa. Nature, 397, 574.
BREMER C.W., DUNGU-KIMBENGA B. & VILJOEN G.J. (1998). Detection of African horsesickness virus in Zebra by RTPCR and the development of different methods for confirming AHSV specificity of RT-PCR products. Proceedings
of the Eighth International Conference on Equine Infectious Diseases, Dubai, 23–26 March 1998. R & W
Publications (Newmarket) Ltd, Newmarket, UK.
COETZER J.A.W. & GUTHRIE.A.J. (2005). African horsesickness. In: Infectious Diseases of Livestock, Second
Edition. Coetzer J.A.W. & Tustin R.C., eds. Oxford University Press, Cape Town, 1231–1246.
ERASMUS B.J. (1976). A new approach to polyvalent immunisation against African horse sickness. In: Proceedings
of the Fourth International Conference on Equine Infectious Diseases, Lyon, France, September 1976. Princeton,
N.J. Veterinary Publications, USA, 401–403.
EUROPEAN COMMISSION (2002). Commission decision of 21 February 2002 amending Annex D to Council Directive
90/426/EEC with regard to the diagnostic tests for African horse sickness. Off. J. European Communities, L53,
37–42.
FERNÁNDEZ J., FERNÁNDEZ-PACHECO P., RODRÍGUEZ B., SOTELO E., ROBLES A., ARIAS M. & SÁNCHEZ-VIZCAÍNO J.M.
(2009). Rapid and sensitive detection of African horse sickness virus by real-time PCR. Res. Vet. Sci., 86, 353–
358.
GRUBMAN M. & LEWIS S. (1992). Identification and characterisation of the structural and non-structural proteins of
African horse sickness virus and determination of the genome coding assignments. Virology, 186, 444–451.
HAMBLIN C., GRAHAM S.D., ANDERSON E.C. & CROWTHER J.R. (1990) A competitive ELISA for the detection of
group-specific antibodies to African horse sickness virus. Epidemiol. Infect., 104, 303–312.
HOUSE C., MIKICIUK P.E. & BERNINGER M.L. (1990). Laboratory diagnosis of African horse sickness: comparison of
serological techniques and evaluation of storage methods of samples for virus isolation. J. Vet. Diagn. Invest., 2,
44–50.
HOUSE J., LOMBARD M., HOUSE C., DUBOURGET P. & MEBUS C. (1992). Efficacy of an inactivated vaccine for African
horse sickness serotype 4. In: Bluetongue, African Horse Sickness and Related Orbiviruses: Proceedings of the
Second International Symposium, Walton T.E. & Osburn B.l., eds. CRC Press, Boca Raton, Florida, USA, 891–
895.
KOEKEMOER M.J.P. (2008). Serotype-specific detection of African horsesickness virus by real-time PCR and the
influence of genetic variations. J. Virol. Methods, 154, 104–110.
KOEKEMOER M.J.P., POTGIETER A.C., PAWESKA J.T. & VAN DIJK A.A. (2000). Development of probes for typing
African horsesickness virus isolates using a complete set of clone VP-2 genes. J. Virol. Methods, 88, 135–144.
LAVIADA M.D., ARIAS M. & SANCHEZ-VIZCAINO J.M. (1993). Characterization of African horse sickness virus serotype
4-induced polypeptides in Vero cells and their reactivity in Western immunoblotting. J. Gen. Virol., 74, 81–87.
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LAVIADA M.D., ROY P. & SANCHEZ-VIZCAINO J.M (1992b). Adaptation and evaluation of an indirect ELISA and
inmunoblotting test for African horse sickness antibody detection. In: Bluetongue, African Horse Sickness and
Related Orbiviruses: Proceedings of the Second International Symposium. Walton T.E. & Osburn B.l., Eds. CRC
Press, Boca Raton, Florida, USA, 646–650.
LAVIADA M.D., SANCHEZ-VIZCAINO J.M., ROY P. & SOBRINO F. (1997). Detection of African horsesickness virus by the
polymerase chain reaction. Invest. Agr. SA., 12, 97–102.
MAREE S. & PAWESKA J.T. (2005). Preparation of recombinant African horse sickness virus VP7 antigen via a
simple method and validation of a VP7-based indirect ELISA for the detection of group-specific IgG antibodies in
horse sera. J. Virol. Methods, 125 (1), 55–65.
MARTINEZ J., DIAZ-LAVIADA M., ROY P., SANCHEZ C., VELA C., SANCHEZ-VIZCAINO J.M. & CASAL I. (1996). Full protection
against AHSV in horses induced by baculovirus-derived AHS virus serotype 4 VP2, VP5 and VP7. J. Gen. Virol., 77,
1211–1221.
MARTINEZ-TORRECUADRADA J., LANGEVELD J., MELOEN R. & CASAL I. (2001). Definition of neutralizing sites on African
horse sickness virus serotype 4 VP2 at the level of peptides. J. Gen. Virol., 82, 2415–2424.
QUAN M., LOURENS C.W., MACLACHLAN N.J., GARDNER I.A. & GUTHRIE A.J. (2010). Development and optimisation of
a duplex real-time reverse transcription quantitative PCR assay targeting the VP7 and NS2 genes of African
horse sickness virus. J. Virol. Methods, 167, 45–52.
RODRIGUEZ-SANCHEZ B., FERNANDEZ-PINERO J., SAILLEAU C., ZIENTARA S., BELAK S., ARIAS M. & SANCHEZ-VIZCAINO J.M.
(2008). Novel gel-based and real-time PCR assays for the improved detection of African horse sickness virus. J. Virol.
Methods, 151, 87–94.
ROY P., HIRASAWA T., FERNANDEZ M., BLINOV V.M. & SANCHEZ-VIZCAINO RODRIGUEZ J.M. (1991). The complete sequence
of the group-specific relationship to bluetongue virus. J. Gen. Virol., 72, 1237–1241.
RUBIO C., CUBILLO M.A., HOOGHUIS H., SANCHEZ-VIZCAINO JM., DIAZ-LAVIADA M., PLATEAU E., ZIENTARA S., CRUCIERE C. &
HAMBLIN C. (1998). Validation of ELISA for the detection of African horse sickness virus antigens and antibodies. Arch.
Virol. (Suppl.), 14, 311–315.
SAILLEAU C., HAMBLIN C., PAWESKA J. & ZIENTARA S. (2000). Identification and differentiation of nine African horse
sickness virus serotypes by RT-PCR amplification of the serotype-specific genome segment 2. J. Gen. Virol., 81, 831–
837.
SAKAMOTO K., PUNYAHOTRA R., MIZUKOSHI N., UEDA S., IMAGAWA H., SUGIURA T., KAMADA M. & FUKUSHO A. (1994).
Rapid detection of African horsesickness virus by the reverse transcriptase polymerase chain reaction (RT-PCR)
using the amplimer for segment 3 (VP3 gene). Arch. Virol., 36 (1–2), 87–97.
SANCHEZ-VIZCAÍNO J.M. (2004). Control and eradication of African horse sickness with vaccine. In: Control of
Infectious Diseases by Vaccination, Schudel A. & Lombard M., eds. Developments in Biologicals, 119, 255–258.
S. Karger AG, Basel, Switzerland.
SCANLEN M., PAWESKA J., VERSCHOOR J. & DIJK A. (2002). The protective efficacy of a recombinant VP2-based African
horsesickness subunit vaccine candidate is determined by adjuvant. Vaccine, 20, 1079–1088.
STONE-MARSCHAT M., CARVILLE A., SKOWRONEK A. & LAEGREID W.W. (1994). Detection of African horse sickness
virus by reverse transcription PCR. J. Clin. Microbiol., 32, 697–700.
TENA C. (2009). Development of serotypes 4 and 9 specific RRT-PCR assays. Annual Meeting of EC National
African Horse Sickness (AHS) Reference Laboratories. Brussels, Belgium, 1 December 2009.
VAN NIEKERT M., VAN STADEN V., VAN DIJK A.A. & HUISMANS H. (2001). Variation of African horsesickness virus
nonstructural protein NS3 in southern Africa. J. Gen. Virol., 82, 149–158.
VENTER M., NAPIER G. & HUISMANS H. (2000). Cloning, sequencing and expression of the gene that encodes the
major neutralisation-specific antigen of Africa horsesickness virus serotype 9. J. Virol. Methods, 86, 41–53.
VERWOERD D.W., HUISMANS H., ERASMUS B.J. (1979). Orbiviruses. In: Comprehensive Virology, Fraenkel-Conrat
H., Wagner R.R., eds. Plenum Press, London, UK, Vol. 14, 285–345.
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VILLALBA R. (2009). Results of the AHS Interlaboratory comparison test 2009: RT-PCR. Annual Meeting of EC
National African Horse Sickness (AHS) Reference Laboratories. Brussels, Belgium, 1 December 2009.
WADE-EVANS A., WOOLHOUSE T., O’HARA R. & HAMBLIN C. (1993). The use of African horse sickness virus VP7
antigen, synthesised in bacteria, and anti-VP7 monoclonal antibodies in a competitive ELISA. J. Virol. Methods,
45, 179–188.
WILLIAMS C.F., INOUE T., LUCUS A.M., ZANOTTO P.M. & ROY Y.P. (1998). The complete sequence of four major
structural proteins of African horse sickness virus serotype 6: evolutionary relationship within and between the
orbivirus. Virus Res., 53, 53–73.
ZIENTARA S., SAILLEAU C., MOULAY S. & CRUCIERE C. (1994). Diagnosis of the African horse sickness virus serotype
4 by a one-tube, one manipulation RT-PCR reaction from infected organs. J. Virol. Methods, 46, 179–188.
*
* *
NB: There are OIE Reference Laboratories for African horse sickness
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for African horse sickness
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2012
CHAPTER 2.5.2.
CONTAGIOUS EQUINE METRITIS
SUMMARY
Definition of the disease: Contagious equine metritis is an inflammatory disease of the proximal
and distal reproductive tract of the mare caused by Taylorella equigenitalis, which usually results in
temporary infertility. It is a nonsystemic infection, the effects of which are restricted to the
reproductive tract of the mare.
When present, clinical signs include endometritis, cervicitis and vaginitis of variable severity and a
slight to copious mucopurulent vaginal discharge. Recovery is uneventful, but prolonged
asymptomatic or symptomatic carriage is established in a proportion of infected mares. Direct
venereal contact during natural mating presents the highest risk for the transmission of
T. equigenitalis from a contaminated stallion or an infected mare. Direct venereal transmission can
also take place by artificial insemination using infective raw, chilled and possibly frozen semen.
Indirectly, infection may be acquired through fomite transmission, manual contamination,
inadequate observance of appropriate biosecurity measures at the time of breeding and at semencollection centres. Stallions can become asymptomatic carriers of T. equigenitalis. The principal
sites of colonisation by the bacterium are the urogenital membranes (urethral fossa, urethral sinus,
terminal urethra and penile sheath). The sites of persistence of T. equigenitalis in the majority of
carrier mares are the clitoral sinuses and fossa and infrequently the uterus. Foals born of carrier
mares may also become carriers. The organism can infect equid species other than horses, e.g.
donkeys.
Identification of the agent: To avoid loss of viability, individual swabs should be fully submerged
in Amies charcoal medium and transported to the testing laboratory under temperature-controlled
conditions for plating out within 48 hours of collection. Growth of T. equigenitalis is likely to take 3–
6 days and may take up to 14 days, but usually does not take longer than 6 days at 37°C on
specialised media in an atmosphere of 5–10% CO2. An incubation time of at least 7 days is
advisable before certifying cultures negative for T. equigenitalis. Identification should include
biochemical characterisation, antigenic testing using specific antibodies and molecular genotyping.
The fastidious nature of T. equigenitalis makes it difficult to isolate and test-breeding of stallions for
detection of the carrier state has been used as a valuable adjunct to cultural examination.
Another species of Taylorella, T. asinigenitalis, has been isolated from male donkeys and horse
mares and stallions in the United States of America and a number of European countries. This
bacterium has not been associated with naturally occurring disease; it has been detected in the
genital tract of male donkeys and would appear to be passed to other donkeys and horses during
mating.
Serological tests: Serology is of value in detecting recent, but not chronic, infection in the mare.
Serum antibody to T. equigenitalis can be detected in mares for 3–7 weeks after infection. It may
also be demonstrated in the occasional carrier mare, but never in the stallion. No individual
serological test described to date has been shown reliably to detect infection. Serological tests can
be used as an adjunct to culture for T. equigenitalis in screening mares recently bred to a carrier
stallion, but must not be used as a substitute for culture.
Requirements for vaccines: Effective vaccines are not yet available.
A. INTRODUCTION
Contagious equine metritis was first described in the United Kingdom (UK) in 1977 (Crowhurst, 1977), after which
it was diagnosed in a number of countries world-wide. It first presented as disease outbreaks characterised by a
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Chapter 2.5.2. — Contagious equine metritis
mucopurulent vaginal discharge originating from inflammation of the endometrium and cervix, resulting in
temporary infertility. The fastidious growth characteristics and slow growth of the causative bacterium, Taylorella
equigenitalis, caused difficulties in initial attempts at culture (Platt et al., 1977), but the disease was reproduced by
experimental challenge with isolated laboratory-grown bacteria (Platt et al., 1978; Swaney & Sahu 1978). Using
appropriate culture conditions, T. equigenitalis can be isolated from infective vaginal discharge. Mares may
experience more than one episode of the disease in a short period of time (Timoney et al., 1977; 1979). Serum
antibody persists for 3–7 weeks after infection, but often it is not detectable for up to 15–21 days after recovery
from acute infection in the mare (Dawson et al., 1978). Most mares recover uneventfully, but some may become
carriers of T. equigenitalis for many months (Platt et al., 1978). Colonisation by T. equigenitalis in the mare is
most consistently demonstrated by swabbing the clitoral fossa and recesses of the clitoral sinuses; it may be
recovered from the cervix and endometrium in pure culture (Platt et al., 1978). Sampling of the clitoral sinuses
should be done with mini-tipped swabs, not the standard type of bacteriological swab. Carriage does not always
adversely affect conception (Timoney et al., 1978), and in such cases pregnancy may proceed to term. The
offspring of such pregnancies may become contaminated during passage through the birth canal, and such foals
may become long-term subclinical carriers (Timoney & Powell, 1982). Many primary cases of infection with
T. equigenitalis in the mare are subclinical, and a frequent indicator of infection is a mare returning in oestrus
prematurely after being bred to a putative carrier stallion.
Carrier mares and stallions act as reservoirs of T. equigenitalis, but stallions, because they mate with numerous
mares, play a much more important role in dissemination of the bacterium. The urogenital membranes of stallions
become contaminated at coitus, leading to a carrier state that may persist for many months or years (Schluter et
al., 1991). Failure to observe appropriate hygienic measures when breeding mares and stallions may also spread
the organism. Other sites of the horse’s body are not known to harbour T. equigenitalis. Most carrier mares are
clitoral carriers of T. equigenitalis. Long-term persistence of the organism in the uterus, though uncommon, can
occur. To detect such carriers of T. equigenitalis, an endometrial or deep cervical swab sample should be taken
routinely in addition to sampling the clitoral area of all mares. Abortion in the mare due to T. equigenitalis is a very
rare occurrence.
Another species of Taylorella, T. asinigenitalis (Jang et al., 2001) has been isolated from male donkeys and horse
mares and stallions in the United States of America (Katz et al., 2000; Meade et al., 2010), and a number of
European countries (Baverud et al., 2006; Breuil et al., 2011; Franco et al., 2009). This bacterium, which has not
been associated with naturally occurring disease, resides in the genital tract of male donkeys and can be passed
to donkeys and horses during mating.
Taylorella equigenitalis is not known to infect humans and it should be handled in the laboratory at biosafety level
2.

Disease control
Prior infection or vaccination are not fully protective (Fernie et al., 1980; Timoney et al., 1979), and failure of
antibody to persist has meant that control of infection has relied entirely on prevention of transmission through the
detection of T. equigenitalis on swabs of the reproductive tract of stallions and mares. In spite of difficulties in
culturing T. equigenitalis, screening mares and stallions prior to and while on the stud farm has successfully
eliminated the disease from thoroughbred horses in countries using a voluntary code of practice. These have
been based on the widely adopted UK’s Horserace Betting Levy Board’s Code of Practice
(www.hblb.org.uk/codes.htm), which is reviewed and updated annually and will reflect current information. The
key recommendations of that Code of Practice are summarised below.
At the start of the breeding season, swabs are taken from all stallions, including those in their first breeding
season, from the urethra, urethral fossa and sinus, penile sheath and pre-ejaculatory fluid, on two occasions no
fewer than 7 days apart. Mares are classified according to the degree of risk that they represent, and the
frequency of sampling is adjusted accordingly. High-risk mares are defined in the Code of Practice as: (a) those
from which T. equigenitalis has been isolated (the high risk status will remain until three sets of negative swabs
have been taken at three different oestrous periods in each of 2 years); (b) mares that have visited any premises
on which T. equigenitalis has been isolated within the previous 12 months; (c) mares arriving from countries
currently listed in the Code of Practice as higher risk; d) all mares that have been in countries that are currently
listed in the Code of Practice as higher risk within the last 12 months.
High risk stallions are defined as: (a) stallions that have not previously been used for breeding purposes;
(b) stallions from which T equigenitalis has been isolated (the ‘high risk’ status will remain until treatment has
been undertaken and required swab results are negative); (c) stallions that have been , on any premises on which
T equigenitalis has been isolated within the last 12 months; (d) stallions that have mated a mare which has not
been swabbed negative in accordance with the Code of Practice. ‘Low risk’ stallions are any stallions not defined
as ‘high risk’. There is a potentially serious problem with contagious equine metritis in this group of horses,
especially those belonging to non-thoroughbred breeds.
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Chapter 2.5.2. — Contagious equine metritis
Results of laboratory tests for T. equigenitalis should be entered on an officially approved certificate, which is sent
to the veterinarians and stallion stud farm managers who supervise the breeding. The certificate should record the
animal’s name, the sites and date of swabbing, the name of the veterinarian taking the swabs, identity of the
testing laboratory, the date the swabs were received and cultured by the laboratory, and whether the swabs were
negative or positive, or whether the culture was overgrown by other bacteria to an extent that the laboratory could
not be confident that small numbers of T. equigenitalis would be detected and, therefore, requiring another set of
swabs to be collected.
Difficulties with the culture of T. equigenitalis due to its fastidious nature necessitate the use of a quality control
system that should be approved before a laboratory is permitted to undertake official testing for contagious equine
metritis and to issue certificates of the test results. The task of quality control should be undertaken by an
experienced, reliable, and impartial microbiology laboratory authorised for the purpose.
Any mares with an abnormal vaginal exudate, or returning to oestrus prematurely, should be investigated and
managed as though infected with T. equigenitalis until results of laboratory testing prove otherwise. Other causes
of endometritis include Pseudomonas aeruginosa, Streptococcus zooepidemicus and certain capsule types of
Klebsiella pneumoniae. Swabs should be examined for these bacteria, and an attempt made to culture and
identify K. pneumoniae and P. aeruginosa so as to establish a differential diagnosis.
If carriers of T. equigenitalis are detected, the organism can be eliminated by treatment with systemic and/or local
antibiotics combined with antiseptic washing of the sites of persistence in the mare and the stallion. Particular
attention should be paid to the clitoral fossa and cleansing the recesses of the clitoral sinuses of mares, where
colonisation by T. equigenitalis is frequently found in carrier animals. A course of treatment may take several
weeks and may need to be repeated before swabbing consistently fails to recover T. equigenitalis from the
treated stallion or mares (Crowhurst et al., 1979). A variable number of carrier mares can be refractive to several
courses of treatment. These may require surgery and ablation of the clitoral sinuses for permanent elimination of
the carrier state in such animals.
Control measures for countries regarded as free from T. equigenitalis infection should be based on the screening
of animals prior to importation and/or during a post-importation quarantine period using swabbing and testing
regimes broadly based on those described above for breeding populations.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent (the prescribed test for international trade)
Various bacteria may be present on the urogenital membranes of horses as harmless commensals and may
interfere with the culture of T. equigenitalis. Some may initially be present in small numbers, but multiply on the
swab before it is cultured depending on the ambient temperature the swab is exposed to in transit to the testing
laboratory. Overgrowth of these organisms on the culture plates may obscure the presence of T. equigenitalis.
Swabs must be placed in a transport medium with activated charcoal, such as Amies medium, to absorb inhibitory
by-products of bacterial metabolism (Swerczek, 1978). The numbers of viable T. equigenitalis decline on swabs
over time, and this effect is more pronounced at higher temperatures (Sahu et al., 1979). Swabs must be kept
cool during transportation and should arrive and be plated out at the laboratory no later than 48 hours after they
were taken. Negative culture results from swabs plated out more than 48 hours after they were taken are
unreliable. Antibiotic treatment for whatever cause should cease at least 7 days (systemic treatment) or 21 days
(local treatment) before swabbing. The presence of antibiotics may sublethally damage T. equigenitalis, which
nonetheless persists on the urogenital membranes but cannot be grown on laboratory media.
Each swab must be inoculated on to 5% (v/v) heated blood, (‘chocolate’), agar plates, produced by heating the
liquid medium, containing blood, at 70–80°C for 12 minutes. When cooled to 45–50°C, trimethoprim (1 µg/ml),
clindamycin (5 µg/ml), and amphotericin B (5–15 µg/ml), is added to the medium (Timoney et al., 1982).
Thymidine, which will inactivate trimethoprim, is present in bacteriological media containing peptone, so it is
important to add 5% lysed horse blood at this stage. Lysed horse blood contains thymidine phosphorylase, which
will inactivate thymidine, thus allowing the trimethoprim to exert its selective effect. This is the preferred medium
for isolating T. equigenitalis; this medium has been used successfully to isolate equally well both streptomycin
resistant and sensitive biotypes of the pathogen and to suppress the growth of many commensal bacteria and
inhibit fungal growth. As inhibiters may prevent the isolation of some strains of T. equigenitalis, swabs should also
be inoculated on to plates of 5% ‘chocolate’ blood agar with a rich peptone agar base containing additional
cysteine (0.83 mM), sodium sulphite (1.59 mM) and a fungicide (5–15 µg/ml amphotericin B). While
T. equigenitalis can be cultivated on blood agar, it will grow better on ‘chocolate’ blood agar as described above.
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Chapter 2.5.2. — Contagious equine metritis
Some manufacturers1 produce a peptone agar base that is quality controlled for its ability to support the growth of
T. equigenitalis. The quality of the commercial agar should be confirmed by the testing laboratory. An important
feature of all good T. equigenitalis media is the absence of fermentable carbohydrates. These do not enhance the
growth of T. equigenitalis, but their fermentation by other bacteria inhibits T. equigenitalis growth (Atherton, 1983;
Fernie et al., 1980). A third medium containing streptomycin sulphate (200 µg/ml) is sometimes used as some
isolates of T. equigenitalis are resistant to this concentration of antibiotic, which serves to reduce the extent of
growth of other bacteria that might otherwise obscure the presence of small numbers of T. equigenitalis
(Swerczek, 1978). However, a streptomycin-sensitive biotype is now the most common strain isolated and will not
be detected on this medium; consequently, it should only be used in conjunction with medium without
streptomycin. Growth by other bacteria, for example Proteus mirabilis, however, may be so extensive that the
laboratory should report that they cannot issue a negative result. In this event, further swabs should be requested
in the hope that the problem will not recur.
Occasionally, the urogenital membranes of stallions or mares will be persistently colonised by another bacterium
that interferes with detection of T. equigenitalis. It may be necessary to attempt to eliminate this by washing and
antibiotic treatment. Swabbing for T. equigenitalis shall not recommence until at least 7 days (systemic treatment)
or 21 days (local treatment) after treatment has stopped. The use of Timoney’s medium (Timoney et al., 1982),
described above, should overcome this difficulty in most cases
All culture media should be subjected to quality control and must support growth of a small inoculum of the
suspect organism before their use on suspect samples. The reference strain of T. equigenitalis must also be
cultured in parallel with the test samples to ensure that the culture conditions are optimal for isolation of this
organism.
The fastidious nature of T. equigenitalis makes it difficult to isolate. Test breeding of stallions has been used to
increase the sensitivity of detection of the carrier state and it has been a valuable adjunct to cultural examination.
The numbers of Taylorella present on the external genitalia of stallions can be very low and may be missed by
culturing alone, but can be detected after multiplication in the mare that has been test bred. The use of test
breeding as an additional diagnostic tool can be especially important in countries that are considered free from
contagious equine metritis.
Plates must be incubated at 35–37°C in 5–10% (v/v) CO2 in air or by use of a candle jar. At least 72 hours is
normally required before colonies of T. equigenitalis become visible, after which time daily inspection is needed.
Rarely, visual detection of colonies may take up to 14 days (Ward et al., 1984). A standard incubation time of at
least 7 days is advisable before certifying cultures negative for T. equigenitalis. Plates should be examined for
contaminants after the first 24 hours’ incubation. Colonies of T. equigenitalis may be up to 2–3 mm in diameter,
smooth with an entire edge, glossy and yellowish grey. Laboratories should be aware that certain countries
require the prolonged incubation period as a standard procedure and should therefore ascertain the particular
import requirements of those countries and/or indicate the incubation period on which their cultural findings are
based.
Taylorella equigenitalis is a Gram-negative, nonmotile, bacillus or cocco-bacillus that is often pleomorphic (up to
6 µm long) and may exhibit bipolar staining. It is catalase positive, phosphatase positive, and strongly oxidase
positive. It is otherwise inert in tests for biochemical activity. If a slow-growing organism is isolated that fits the
description for cellular morphology and that is strongly oxidase positive, it should be tested for reactivity with T.equigenitalis-specific antiserum.
A variety of serotyping tests have been developed ranging in complexity from slide agglutination to direct or
indirect immunofluorescence. Each method has its advantages and disadvantages. The disadvantage of the slide
agglutination test is that occasionally autoagglutination of isolates occurs: culturing in bottled CO2 in air, as
opposed to in a candle jar, may reduce autoagglutination (Ter Laak & Wagenaars, 1990). It has been suggested
that immunofluorescence can be used to identify autoagglutinating isolates, some workers have reported crossreaction with Mannheimia haemolytica but this is very rare. If a cross-reaction is suspected, it may be necessary
to repeat the test using adsorbed antisera (Ter Laak & Wagenaars, 1990). The specificity of the
immunofluorescence test can be improved by the use of monoclonal antibodies that are now available.
Nevertheless polyclonal and monoclonal antibodies can be used in the differentiation of T. equigenitalis and
T. asinigenitalis (Breuil et al., 2010) 2.
Antiserum is produced by vaccinating rabbits with killed T. equigenitalis. A number of different immunisation
regimes can be employed, ranging from those used for producing Escherichia-coli-typing antisera to immunisation
together with an adjuvant, such as Freund’s incomplete. Monoclonal antibodies are available commercially that
1
2
834
For example, Mast Diagnostics, Mast House, Derby Road, Bootle, Merseyside L20 1EA, United Kingdom (UK), and
Lab M, Tomley House, Wash Lane, Bury BL9 6AU, UK.
Institut Pourquier, 326 rue de la Galera, Parc Euromedicine, 34090 Montpelier, France. E-mail: [email protected]
OIE Terrestrial Manual 2012
Chapter 2.5.2. — Contagious equine metritis
provide a highly specific means of identifying T. equigenitalis. A standard strain, such as NCTC 111843, should be
used for immunisation. However, the most important consideration is the specificity of the antiserum produced. It
should agglutinate T. equigenitalis, but fail to agglutinate other bacteria that might be cultured from horse
urogenital membranes, even if rarely. In particular, it should not agglutinate any oxidase-positive and Gramnegative rods, such as Mannheimia haemolytica, Actinobacillus equuli, Bordetella bronchiseptica (to which
T. equigenitalis is closely related, see Bleumink-Pluym et al. (1993), Oligella urethralis and Pseudomonas
aeruginosa. Taylorella asinigenitalis has similar, though not identical, colonial appearance and cultural
characteristics and gives identical biochemical test results to those used to confirm the identity of T. equigenitalis.
There is even serological cross-reactivity between the two organisms. Differentiation of T. asinigenitalis from
T. equigenitalis is possible using the polymerase chain reaction (PCR) or 16S rDNA sequencing and biochemical
reactivity (Baverud et al., 2006; Breuil et al., 2011; Duquesne et al., 2007; Wakeley et al., 2006).
A latex agglutination kit is available commercially for the antigenic identification of T. equigenitalis. It is based on
polyclonal antibodies produced using methods similar to those described above. This is widely used by routine
testing laboratories for the confirmation of the identity of colonies growing on selective medium that give a
biochemical reaction consistent with T. equigenitalis. As T. equigenitalis is antigenically relatively distinct, and
small amounts of cross-reactive antibody are easily absorbed during production of the reagent, the test has
proved to be highly specific and sensitive. It should be emphasised that it will not necessarily distinguish strains of
T. equigenitalis from T. asinigenitalis.
Molecular testing methods such as PCR and real-time PCR have been applied to the detection of
T.equigenitalis both directly (using swabs taken from sampling sites) and indirectly (from cultures grown from
swabs) (Bleumink-Pluym et al., 1994). In studies carried out in the UK the PCR was shown to be highly specific
and was able to detect very small numbers of T. equigenitalis in the presence of very large numbers of bacteria
comprising the background flora harvested from culture plates inoculated with samples from the equine urogenital
tract. In Japan the field application of the PCR was evaluated for the eradication of contagious equine metritis. It
was demonstrated that the PCR was more sensitive than culture for the detection of T. equigenitalis from genital
swabs of horses (Anzai et al., 2002; Moore et al., 2001). A real-time PCR was developed in the UK for use
directly on genital swabs and compared with culture (Wakeley et al., 2006) and this has been subsequently used
for pre-breeding screening studies (Ousey et al., 2009). There was no significant difference in the performance of
the direct PCR and culture, but the PCR had the added advantage of speed of result and also differentiated
T. equigenitalis from T. asinigenitalis. Commercial PCR kits are available for the detection of T. equigenitalis and
these may be used to enhance the testing capabilities of authorised laboratories.
2.
Serological tests
No serological test described to date will, by itself, reliably detect infection for diagnosis and control. However, the
complement fixation test has been used successfully as an adjunct to culture for T. equigenitalis in screening
mares between 21 and 45 days after being bred to a suspect carrier stallion.
C. REQUIREMENTS FOR VACCINES
Effective vaccines that protect against contagious equine metritis or prevent colonisation by T. equigenitalis are
not yet available.
REFERENCES
ANZAI T., WADA R., OKUDA T. & AOKI T. (2002). Evaluation of the field application of PCR in the eradication of
contagious equine metritis from Japan. J. Vet. Med. Sci., 64, 999–1002.
ATHERTON J.G. (1983). Evaluation of selective supplements used in media for the isolation of the causative
organism of contagious equine metritis. Vet. Rec., 113, 299–300.
BAVERUD V., NYSTROM C. & JOHANSSON K.-E. (2006). Isolation and identification of Taylorella asinigenitalis from the
genital tract of a stallion, first case of a natural infection. Vet. Microbiol., 116, 294–300.
BLEUMINK-PLUYM N.M.C., VAN DIJK L., VAN VLIET A.H., VAN DER GIESSEN J.W. & VAN DER ZEIJST B.A. (1993).
Phylogenetic position of Taylorella equigenitalis determined by analysis of amplified 16S ribosomal DNA
sequences. Int. J. Syst. Bacteriol., 43, 618–621.
3
Obtainable from the National Collection of Type Cultures, Colindale, London, UK.
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Chapter 2.5.2. — Contagious equine metritis
BLEUMINK-PLUYM N.M.C., WERDLER M.E.B., HOUWERS D.J., PARLEVLIET J.M., COLENBRANDER B. & VAN DER ZEIJST
B.A.M. (1994). Development and evaluation of PCR test for detection of Taylorella equigenitalis. J. Clin.
Microbiol., 32, 893–896.
BREUIL M.S.F., DUQUESNE F., SEVIN C., LAUGIER C. & PETRY S. (2010). Indirect immunofluorescence test using
polyclonal antibodies for the detection of T. equigenitalis. Res. Vet. Sci., 88, 369–371.
BREUIL M.F., DUQUESNE F., LAUGIER C. & PETRY S. (2011a). Phenotypic and 16S ribosomal RNA gene diversity of
Taylorella asinigenitalis strains isolated between 1995 and 2008. Vet. Microbiol., 148, 260–266.
CROWHURST R.C. (1977). Genital infection in mares. Vet. Rec., 100, 476.
CROWHURST R.C., SIMPSON D.J., GREENWOOD R.E.S. & ELLIS D.R. (1979). Contagious equine metritis. Vet. Rec.,
104, 465.
DAWSON F.L.M., BENSEN J.A. & CROXTON-SMITH P. (1978). The course of serum antibody development in two
ponies experimentally infected with contagious metritis. Equine Vet. J., 10, 145–147.
DUQUESNE F., PRONOST S., LAUGIER C. & PETRY S. (2007). Identification of Taylorella equigenitalis responsible for
contagious equine metritis in equine genital swabs by direct polyermase chain reaction. Res. Vet. Sci., 82, 47–49.
FERNIE, BATTY I., WALKER P.D., PLATT H., MACKINTOSH M.E. & SIMPSON D.J. (1980). Observations on vaccine and
post-infection immunity in contagious equine metritis. Res. Vet. Sci., 28, 362–367.
FRANCO A., DONATI V., TROIANO P., LORENZETTI R., ZINI M., AUTORINO G.L., PETRELLA A., MAGGI A. & BATTISTI A.
(2009). Detection of Taylorella asinigenitalis in donkey jacks in Italy. Vet. Rec., 165, 540–541.
JANG S.S., DONAHUE J.M., ARATA A.B., GORIS J., HANSEN L.M., EARLEY D.L., VANDAMME P.A., TIMONEY P.J. & HIRSH
D.C. (2001). Taylorella asinigenitalis sp. nov., a bacterium isolated from the genital tract of male donkeys (Equus
asinus). Int. J. Syst. Evol. Microbiol., 51, 971–976.
KATZ J.B., EVANS L.E., HUTTO D.L., SCHROEDER-TUCKER L.C., CAREW A.M., DONAHUE J.M. & HIRSH D.C. (2000).
Clinical, bacteriologic, serologic and pathologic features of infections with atypical Taylorella equigenitalis in
mares. J. Am. Vet. Med. Assoc., 216 (12), 1945–1948.
MEADE B.J., TIMONEY P.J., DONAHUE J.M., BRANSCUM A.J., FORD R. & ROWE R. (2010). Initial occurrence of
Taylorella asinigenitalis and its detection in nurse mares, a stallion and donkeys in Kentucky. Prev. Vet. Med., 95,
292–296.
MOORE J.E., BUCKLEY T.C., MILLAR B.C., GIBSON P., CANNON G., EGAN C., COSGROVE H., STANBRIDGE S., ANZAI T.,
MATSUDA M. & MURPHY P.G. (2001). Molecular surveillance of the incidence of Taylorella equigenitalis and
Pseudomonas aeruginosa from horses in Ireland by sequence-specific PCR. Equine Vet. J., 33, 319–322.
OUSEY J.C., PALMER L., CASH R.S.G., GRIMES K.J., FLETCHER A.P., BARRELET A., FOOTE A.K., MANNING F.M. &
RICKETTS S.W. (2009) An investigation into the suitability of a commercial real-time PCR assay to screen for
Taylorella equigenitalis in routine prebreeding equine genital swabs. Equine Vet. J., 41 (9), 878–882.
PLATT H., ATHERTON J.G., DAWSON F.L.M. & DURRANT D.S. (1978). Developments in contagious equine metritis.
Vet. Rec., 102, 19.
PLATT H., ATHERTON J.G., SIMPSON D.J., TAYLOR C.E.D., ROSENTHAL R.O., BROWN D.F.J. & WREGHITT T.G. (1977).
Genital infection in mares. Vet. Rec., 101, 20
SAHU S.P., DARDIRI A.H., ROMMEL F.A. & PIERSON R.E. (1979). Survival of contagious equine metritis bacteria in
transport media. Am. J. Vet. Res., 40, 1040–1042
SCHLUTER H., KULLER H., FREIDREICH U., SELBITZ H., MARWITZ T., BEYER C. & ULLRICH E. (1991). Epizootiology and
treatment of contagious equine metritis (CEM), with particular reference to the treatment of infected stallions.
Prakt. Tierarzt., 72, 503–511.
SWANEY L.M. & SAHU S.P. (1978). CEM: bacteriological methods. Vet. Rec., 102, 43.
SWERCZEK T.W. (1978). Inhibition of the CEM organism by the normal flora of the reproductive tract. Vet. Rec.,
103, 125.
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TER LAAK E.A. & WAGENAARS C.M.F. (1990). Autoagglutination and the specificity of the indirect fluorescent
antibody test applied to the identification of Taylorella equigenitalis. Res. Vet. Sci., 49, 117–119
TIMONEY P.J., O’REILLY P.J., MCARDLE J.F., WARD J. & HARRINGTON A.M. (1979). Responses of mares to
rechallenge with the organism of contagious equine metritis 1977. Vet. Rec., 104, 264.
TIMONEY P.J. & POWELL D.G. (1982). Isolation of the contagious equine metritis organism from colts and fillies in
the United Kingdom and Ireland. Vet. Rec., 111, 478–482.
TIMONEY P.J., SHIN S.J. & JACOBSON R.H. (1982). Improved selective medium for isolation of the contagious equine
metritis organism. Vet. Rec., 111,107–108.
TIMONEY P.J., WARD J. & KELLY P. (1977). A contagious genital infection of mares. Vet. Rec., 101, 103
TIMONEY P.J., WARD J. & MCARDLE J.F. (1978). CEM and the foaling mare. Vet. Rec., 102, 246–247
WAKELEY P.R., ERRINGTON J., HANNON S., ROEST H.I.J., CARSON T. HUNT B. & HEATH P. (2006). Development of a
real time PCR for the detection of Taylorella equigenitalis directly from genital swabs and discrimination from
T. asinigenitalis. Vet. Microbiol., 118, 247–254.
WARD J., HOURIGAN M., MCGUIRK J. & GOGARTY A. (1984). Incubation times for primary isolation of contagious
equine metritis organism. Vet. Rec., 114, 298.
*
* *
NB: There are OIE Reference Laboratories for Contagious equine metritis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for Contagious equine metritis
OIE Terrestrial Manual 2012
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.3.
DOURINE
SUMMARY
Dourine is a chronic or acute contagious disease of breeding solipeds that is transmitted directly
from animal to animal during coitus. The causal organism is Trypanosoma equiperdum (Doflein,
1901).
Dourine is the only trypanosomosis that is not transmitted by an invertebrate vector. Trypanosoma
equiperdum differs from other trypanosomes in that it is primarily a tissue parasite that rarely
invades the blood. There is no known natural reservoir of the parasite other than infected equids. It
is present in the genital secretions of both infected males and females. The incubation period,
severity, and duration of the disease vary considerably; it is often fatal, but spontaneous recoveries
do occur but latent carriers do exist. Subclinical infections occur, and donkeys and mules are more
resistant than horses and may remain inapparent carriers. Infection is not always transmitted by an
infected animal at every copulation. Rats can be infected experimentally, and can be used to
maintain strains of the parasite indefinitely. Trypanosoma equiperdum strains are best stored in
liquid nitrogen.
The clinical signs are marked by periodic exacerbation and relapse, ending in death, sometimes
after paraplegia or, possibly, recovery. Fever, local oedema of the genitalia and mammary glands,
cutaneous eruptions, incoordination, facial paralysis, ocular lesions, anaemia, and emaciation may
all be observed. Oedematous cutaneous plaques, 5–8 cm in diameter and 1 cm thick, are
pathognomonic.
Identification of the agent: Definitive diagnosis depends on the recognition of clinical signs and
identification of the parasite. As this is rarely possible, diagnosis is usually based on clinical signs,
together with serological evidence from complement fixation (CF) tests.
Serological tests: Humoral antibodies are present in infected animals whether or not they display
clinical signs. The CF test is used to confirm infection in clinical cases or in latent carriers.
Noninfected animals, especially donkeys, often yield unclear results. The indirect fluorescent
antibody test can be used to confirm infection or resolve inconclusive CF test results. Enzymelinked immunosorbent assays are also used.
Requirements for vaccines and diagnostic biologicals: There are no vaccines available. The
only effective control is through the slaughter of infected animals. Good hygiene is essential during
assisted matings because infection may be transmitted through contaminated fomites.
A. INTRODUCTION
Dourine is a chronic or acute contagious disease of breeding solipeds that is transmitted directly from animal to
animal during coitus. The causal organism is Trypanosoma equiperdum (Doflein, 1901). Dourine is also known
under other names: mal de coït, el dourin, morbo coitale maligno, Beschälseuche, slapsiekte, sluchnaya bolyezn,
and covering disease (Barner, 1963; Hoare, 1972).
Although the disease has been known since ancient times, its nature was established only in 1896 when Rouget
discovered trypanosomes in infected Algerian horses. Dourine is the only trypanosomosis that is not transmitted
by an invertebrate vector. Trypanosoma equiperdum differs from other trypanosomes in that it is primarily a tissue
parasite that is rarely detected in the blood. There is no known natural reservoir of the parasite other than infected
equids.
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Chapter 2.5.3. — Dourine
Infection is transmitted during copulation, more commonly from stallion to mare, but also from mare to stallion,
due to the presence of the parasite in the seminal fluid and mucous exudate of the penis and sheath of the
infected male, and in the vaginal mucus of the infected female. Initially, parasites are found free on the surface of
the mucosa or between the epithelial cells of a newly infected animal. Invasion of the tissues takes place, and
oedematous patches appear in the genital tract. Parasites then may pass into the blood, where they are carried to
other parts of the body. In typical cases, this metastatic invasion gives rise to characteristic cutaneous plaques.
The incubation period, severity and duration of the disease vary considerably. In South Africa, the disease is
typically chronic, usually mild, and may persist for several years (Henning, 1955). In other areas, such as northern
Africa and South America, the disease tends to be more acute, often lasting only 1–2 months or, exceptionally,
1 week. Although dourine is a fatal disease with an average mortality of 50% (especially in stallions), spontaneous
recovery can occur. Subclinical infections are recognised. Donkeys and mules are more resistant than horses.
As trypanosomes are not continually present in the genital tract throughout the course of the disease,
transmission of the infection does not necessarily take place at every copulation involving an infected animal.
Transmission of infection from mare to foal can occur via the mucosa, such as the conjunctiva. Mares’ milk has
been shown to be infectious. Animals other than equids can be infected experimentally. Rat-adapted strains can
be maintained indefinitely; infected rat blood can be satisfactorily cryopreserved. Antigens for serological tests are
commonly produced from infected laboratory rats.
The disease is marked by stages of exacerbation, tolerance or relapse, which vary in duration and which may
occur once or several times before death or recovery. The signs most frequently noted are: pyrexia, tumefaction
and local oedema of the genitalia and mammary glands, oedematous cutaneous eruptions, knuckling of the joints,
incoordination, facial paralysis, ocular lesions, anaemia, and emaciation. A pathognomonic sign is the
oedematous plaque consisting of an elevated lesion in the skin, up to 5–8 cm in diameter and 1 cm thick. The
plaques usually appear over the ribs, although they may occur anywhere on the body, and usually persist for
between 3 and 7 days. They are not a constant feature.
Generally, the oedema disappears and returns at irregular intervals. During each recess, an increasing extent of
permanently thickened and indurated tissue can be seen. The vaginal mucosa may show raised and thickened
semitransparent patches. Folds of swollen membrane may protrude through the vulva. It is not uncommon to find
oedema of the mammary glands and adjacent tissues. Depigmentation of the genital area, perineum, and udder
may occur. In the stallion, the first clinical sign is a variable swelling involving the glans penis and prepuce. The
oedema extends posteriorly to the scrotum, inguinal lymph nodes, and perineum, with an anterior extension along
the inferior abdomen. In stallions of heavy breeds, the oedema may extend over the whole floor of the abdomen.
Pyrexia is intermittent; nervous signs include incoordination, mainly of the hind limbs, lips, nostrils, ears, and
throat. Facial paralysis is usually unilateral. In fatal cases, the disease is usually slow and progressive, with
increasing anaemia and emaciation, although the appetite remains good almost throughout.
At post-mortem examination, gelatinous exudates are present under the skin. In the stallion, the scrotum, sheath,
and testicular tunica are thickened and infiltrated. In some cases the testes are embedded in a tough mass of
sclerotic tissue and may be unrecognisable. In the mare, the vulva, vaginal mucosa, uterus, bladder, and
mammary glands may be thickened with gelatinous infiltration. The lymph nodes, particularly in the abdominal
cavity, are hypertrophied, softened and, in some cases, haemorrhagic. The spinal cord of animals with paraplegia
is often soft, pulpy and discoloured, particularly in the lumbar and sacral regions.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
A definitive diagnosis depends on the recognition of the clinical signs and the demonstration of the parasite. This
is rarely possible because: (a) although the clinical signs and gross lesions in the developed disease may be
pathognomonic, they cannot always be identified with certainty, especially in the early stages or in latent cases;
they can be confused with other conditions, such as coital exanthema (moreover, in some countries [e.g. in South
America], T. evansi infections give rise to similar clinical signs); (b) the trypanosomes are only sparsely present
and are extremely difficult to find, even in oedematous areas; and (c) the trypanosomes are only fleetingly present
in the blood, and in small numbers that defy detection. For unknown reasons, no parasite strain of T. equiperdum
has been isolated in any country of the world since 1982 and most of the strains currently available in national
veterinary diagnostic laboratories are related to T. evansi (Claes et al., 2003).
In practice, diagnosis is based on clinical evidence supported by serology. Recently, other approaches have been
studied and reported on (Claes et al., 2003).
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Chapter 2.5.3. — Dourine
In infected animals, trypanosomes are present, in low numbers only, in lymph and oedematous fluids of the
external genitalia, in the vaginal mucus, and fluid contents of plaques. They are usually undetectable in the blood,
but may be found in the urethral or vaginal mucus collected from preputial or vaginal washings or scrapings 4–
5 days after infection. Later, parasites may be found in the fluid contents of oedemas and plaques, especially
shortly after their eruption. The skin of the area over the plaque should be washed, shaved and dried, and the
fluid contents aspirated by syringe. Blood vessels should be avoided. The fresh aspirate is examined
microscopically for motile trypanosomes. These are present for a few days only, so that lesions should be
examined at intervals. The parasite is rarely found in thick blood films, but is sometimes detectable after
centrifuging blood and examining the recentrifuged plasma.
As dourine is the only trypanosome to affect horses in temperate climates, the observation of trypanosomes in
thick blood films is sufficient for a positive diagnosis. In countries where nagana or surra occur, it is difficult to
distinguish T. equiperdum microscopically (morphology, motility) from other members of the subgenus
Trypanozoon (T. evansi, T. brucei). In particular, T. equiperdum and T. evansi cannot be differentiated on the
basis of morphological criteria. Both are monomorphic, slender trypomastigotes with a free flagellum, although
pleomorphic, stumpy, proteonuclear forms are recognised. However, in kinetopastic strains, the presence of maxicircles in T. equperdum and the absence in T. evansi provides a possible differentiation (Wassal et al., 1991).
Typical strains of the parasite range in length from 15.6 to 31.3 µm.
2.
Serological tests
Humoral antibodies are present in infected animals, whether they display clinical signs or not. The complement
fixation (CF) test (UK Ministry of Agriculture, Fisheries and Food [MAFF], 1986) is used to confirm clinical
evidence and to detect latent infections. Uninfected equids, particularly donkeys and mules, often give
inconsistent or nonspecific reactions because of the anticomplementary effects of their sera. In the case of
anticomplementary sera, the indirect fluorescent antibody (IFA) test is of advantage. There is no internationally
adopted protocol. Cross-reactions are possible due to the presence in some countries of other trypanosomes, for
example, T. cruzi and T. evansi. Enzyme-linked immunosorbent assays (ELISAs) are also used. Trypanosoma
equiperdum is closely related to other Old World trypanosomes, including T. brucei and T. evansi. Members of
this genus all share conserved cytoskeletal elements that provoke a strong and cross-reactive serological
response. All diagnostic antigens and antisera (monoclonal and polyclonal) currently available for use in
serodiagnostic testing contain these conserved elements or antibodies to them, and therefore none of the
serological procedures described below is specific for dourine. The diagnosis of dourine must include history,
clinical, and pathological findings as well as serology. Significant improvements in dourine serodiagnosis will
require development of more trypanosome-specific subunit antigens and antibodies to them.
a)
Complement fixation test (the prescribed test for international trade)
Standard or microplate techniques may be used (Herr et al., 1985). Guinea-pig serum (available
commercially) is used as a source of complement. Other reagents are sheep red blood cells (RBCs) washed
in veronal buffer, and rabbit haemolytic serum (i.e. rabbit anti-sheep RBC) (commercial) as well as known
negative and positive control sera.
840

Antigen production
i)
A rat is inoculated with T. equiperdum cryopreserved stock. The rat must be free from T. lewisi, which
could be achieved by injection with neoarsphenamine, but is better accomplished by using specificpathogen free rats. Adult rats receive 0.5–1.0 ml of rapidly thawed frozen stabilate, intramuscularly or
intraperitoneally. At maximum parasitaemia, blood is collected into an anticoagulant, such as heparin,
which will serve as a stock culture for the inoculation of additional rats.
ii)
Twenty large rats are inoculated intramuscularly or intraperitoneally with 0.5–1.0 ml of this stock
culture. All rats are to have a heavy infection concurrently. If necessary, the dose is adjusted and
additional rats are inoculated to reach maximum parasitaemia at the desired time of 72–96 hours. Rats
usually die within 3–5 days; prior to this, blood is taken from the tail for thick smears and examined
microscopically. When parasitaemia is maximal, the rat is killed and blood is collected in Alsevers or
acid–citrate–dextrose (ACD) saline solution. If parasitaemia is not synchronous, blood can be collected
and held in Alsevers or ACD saline at 4°C until blood has been collected from all the rats.
iii)
The blood is filtered through muslin gauze and centrifuged at 800 g for 4 minutes. The RBCs are
mostly deposited while the trypanosomes remain in suspension.
iv)
The supernatant fluid is transferred to a fresh tube; the upper layer of RBCs is mixed with
trypanosomes to a second tube, and the next layer to a third. Alsevers or ACD saline is added to tubes
2 and 3 to prevent clotting of cells. All tubes are mixed and centrifuged at 1500 g for 5 minutes.
OIE Terrestrial Manual 2012
Chapter 2.5.3. — Dourine
v)
The supernatant fluid is discarded and the upper white layer of trypanosomes is transferred from all
tubes into a clean tube. The next pink layer is transferred into a second tube, and the lower layer to a
third tube.
vi)
Physiological saline is added and mixed and the tubes are centrifuged again at 1500 g for 5 minutes to
separate the trypanosomes. The washing step is repeated until all the trypanosomes are collected as a
pure white mass. Ten rats should produce 3–5 g of antigen. This purification procedure can also be
carried out using a column of DEAE (diethylaminoethyl) cellulose in a solution of phosphate buffered
saline (PBS) containing glucose, pH 8.0 (Lanham & Godfrey, 1970).
vii)
The concentrated trypanosomes are diluted with two volumes of veronal buffer and 5%
polyvinylpyrrolidone as a cryopreservative. Before use in CF tests, the antigen must be dispersed to a
fine suspension with a hand-held or motorised ground glass homogeniser chilled in ice (Watson, 1920).
This antigen may be divided into aliquots, frozen and lyophilised.
The antigen is standardised by titration against a 1/5 dilution of a standard low-titre antiserum.
Sera: Positive and negative sera should be inactivated at 58°C for 30 minutes before being used in the tests.
Mule and donkey sera are normally inactivated at 62°C for 30 minutes. The USDA complement fixation
protocol calls for inactivation of sera for 35 minutes (United States Department of Agriculture [USDA], 2006).
Dilutions of sera that are positive in the screening test are titrated against two units of antigen. Test sera are
screened at a dilution of 1/5. Sera showing more than 50% complement fixation at this dilution are usually
deemed to be positive.
Anticomplementary sera: If the anticomplementary control shows only a trace, this may be ignored. For all
other anticomplementary sera, the activity must be titrated. A duplicate series of dilutions is made and the
sample is retested using T. equiperdum antigen in the first row and veronal buffer only in the second. The
second row gives the titre of the anticomplementary reaction. Provided the first row shows an end-point that
is at least three dilutions greater than the second, the anticomplementary effect may be ignored and the
sample rated as positive. If the results are any closer, a fresh sample of serum must be requested. Dilution
of the serum 1/2 and heat inactivation at 60–63°C for 30 minutes may result in reduction or removal of the
anticomplementary effect.
Buffers and reagents: 0.15 M veronal buffered saline, pH 7.4, is used for diluting reagents and for washing
sheep RBCs. Antigen is pretested by checkerboard titration, and two units are used in the test. Guinea-pig
complement (C) is tested for its haemolytic activity, and diluted to provide two units for the test. Sheep RBCs
in Alsever’s or ACD saline solution are washed three times. A 3% solution is used for the haemolytic system.
The USDA protocol calls for 2% solution in the microtitration procedure with confirmation in a tube test with
3% RBC (USDA, 2006). Titrated rabbit-anti-sheep RBCs – the rabbit haemolytic serum – is taken at double
the concentration of its haemolytic titre (two units). All test sera, including positive and negative control sera,
are inactivated at a 1/5 dilution before testing.
•
Primary dilutions
i)
100 µl of test serum is diluted with 400 µl of veronal buffer (1/5).
ii)
100 µl of both positive and negative control sera is diluted with 400 µl of veronal buffer (1/5).
iii)
The solutions are incubated in a water bath at 58°C for 30 minutes to inactivate complement and
destroy anticomplementary factors.
•
Screening test procedure
i)
25 µl of inactivated test serum is placed in each of three wells.
ii)
25 µl of inactivated control serum is placed in each of three wells.
iii)
25 µl of T. equiperdum antigen diluted to contain two units is placed in the first well only for each
serum.
iv)
25 µl of complement diluted to contain two units is added to the first two wells only for each serum.
v)
25 µl veronal buffer, pH 7.4, is added to the second well for each serum (anticomplementary well).
vi)
50 µl veronal buffer, pH 7.4, is added to the third well for each serum (lysis activity well).
vii)
The complement control is prepared.
viii) The plate is shaken on a microshaker sufficiently to mix the reagents.
ix)
The plate is incubated for 1 hour in a water bath, incubator or in a humid chamber at 37°C.
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x)
The haemolytic system is prepared. After the first 50 minutes of incubation, the sheep RBCs are
sensitised by mixing equal volumes of rabbit haemolytic serum, diluted to contain two units per 50 µl,
and a 3% suspension of washed RBCs; the solution is mixed well and incubated for 10 minutes at
37°C.
xi)
After incubation, 50 µl of haemolytic system is added to each well.
xii)
The plate is shaken on a microshaker sufficiently to mix the reagents.
xiii) The plate is incubated for 30 minutes at 37°C. To aid in reading the results, the plates can be
centrifuged after incubation.
xiv) Reading the results: the plate is viewed from above with a light source beneath it. The fixation in every
well is assessed by estimating the proportion of cells not lysed. The degree of fixation is expressed as
0, 1+, 2+, 3+, 4+ (0%, 25%, 50%, 75% or 100% cells not lysed). Reactions are interpreted as follows:
4+, 3+, 2+ = positive, 1+ = suspicious, trace = negative, complete haemolysis = negative.
xv)
b)
End-point titration: All sera with positive reactions at 1/5 are serially double diluted and tested
according to the above procedure for end-point titration.
Indirect fluorescent antibody test
An IFA test for dourine can also be used (MAFF, 1986) as a confirmatory test or to resolve inconclusive
results obtained by the CF test. The test is performed as follows:
Antigen: (For method, see preparation of CF test antigen in Section B.2.a) Blood is collected into
heparinised vacutainers or into a solution of acid–citrate–dextrose from an animal in which the number of
trypanosomes is still increasing (about ten parasites per 10× microscope field should be present).
i)
The blood is centrifuged for 10 minutes at 800 g.
ii)
One to two volumes of PBS is added to the packed RBCs, the mixture is agitated, and smears are
made that evenly cover the whole slide.
iii)
The smears are air-dried and then wrapped in bundles of four, with paper separating each slide. The
bundles of slides are wrapped in aluminium foil, sealed in an airtight container over silica gel, and
stored at –20°C or –76°C.
iv)
Slides stored at –20°C should retain their activity for about 1 year, at –76°C they should remain
useable for longer.
Acid–citrate–dextrose solution: Use 15 ml per 100 ml of blood.
Conjugate: Fluorescein-labelled sheep anti-horse immunoglobulins.
•
Test procedure
i)
The antigen slides are allowed to reach room temperature in a desiccator. An alternative method is to
remove slides directly from the freezer and fix them in acetone for 15 minutes.
ii)
The slides are marked out.
iii)
Separate spots of test sera diluted in PBS are applied, and the slides are incubated in a humid
chamber in a water bath at 37°C for 30 minutes.
iv)
The slides are washed in PBS, pH 7.2, three times for 5 minutes each, and air-dried.
v)
Fluorescein-labelled conjugate is added at the correct dilution. Individual batches of antigen and
conjugate should be titrated against each other using control sera to optimise the conjugate dilution.
The slides are incubated in a humid chamber in a water bath at 37°C for 30 minutes.
vi)
The slides are washed in PBS, three times for 5 minutes each, and air-dried. An alternative method, to
reduce background fluorescence, is to counter-stain, using Evans Blue (0.01% in distilled water) for
1 minute, rinse in PBS and then air dry
vii)
The slides are mounted in glycerol/PBS (50/50) or immersion oil (commercially available, nonfluorescing grade).
viii) The slides are then examined under UV illumination. Incident light illumination is used with barrier filter
K 530 and exciter filter BG 12. Slides may be stored at 4°C for 4–5 days. Sera diluted at 1/80 and
above showing strong fluorescence of the parasites are usually considered to be positive. Estimating
the intensity of fluorescence demands experience on the part of the observer.
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Chapter 2.5.3. — Dourine
Standard positive and negative control sera should be included in each batch of tests, and due consideration
should be given to the pattern of fluorescence in these controls when assessing the results of test sera.
c)
Enzyme-linked immunosorbent assay
The ELISA has been developed and compared with other serological tests for dourine (Bundesinstitut für
gesundheitlichen Verbraucherschutz und Veterinärmedizin, 1995; Wassall et al., 1991).
Carbonate buffer, pH 9.6, for antigen coating on to microtitre plates: Na2CO3 (1.59 g); NaHCO3 (2.93g); and
distilled water (1 litre).
Blocking buffer: Carbonate buffer + 3% fetal calf serum (FCS).
PBS, pH 7.4, with Tween 20 (PBST) for washing: KH2PO4 (0.2 g); Na2HPO4 × 12 H2O (2.94 g); NaCl
(8.0 g); KCl (0.2 g in 1 litre distilled water), and Tween 20 (0.5 ml).
Sample and conjugate buffer: PBST + 6% FCS.
Citric phosphate buffer: Citric acid monohydrate (4.2 g in 200 ml distilled water); Na2HPO4 × 12 H2O (in
200 ml distilled water). Both components are mixed at equal volumes.
Substrate indicator system: ABTS (2,2’-Azino-bis-[3-ethylbenzothiazoline-6-sulphonic acid]) (40 mg) is
dissolved in citric phosphate buffer (100 ml), and stored at 4°C in the dark. Just before use, 100 µl of
1/40 H2O2 is added to 10 ml of ABTS.
Conjugate: Rabbit anti-horse IgG (H+L) PO or IgY anti-horse Ig-PO.
Antigen: Lyophilised T. equiperdum antigen (0.5 ml) is reconstituted with coating buffer (5 ml), sonicated
twice for 10 seconds each at 12 µm peak to peak, and centrifuged at 10,000 g for 4 minutes. The
supernatant is further diluted to a pretested working dilution (e.g. 1/500).
•
Test procedure
i)
Wells in columns 2, 4, 6, etc., are charged with 50 µl of antigen, columns 1, 3, 5, etc., are charged with
the same amount of carbonate buffer. The plate is incubated for 40 minutes at 37°C in a humid
chamber, washed in tap water, and 50 µl of blocking buffer is added to each well. The plate is
incubated for 20 minutes, washed in tap water followed by three wash cycles with PBST, with soaking
times of 3 minutes/cycle.
ii)
50 µl of test samples and equine control sera prediluted 1/100 in sample/conjugate buffer is added in
parallel to wells with and without antigen. The plate is incubated for 30 minutes, washed in tap water,
followed by three wash cycles with PBST.
iii)
Properly diluted conjugate in sample/conjugate buffer is added in volumes of 50 µl to all wells. The
plate is incubated for 30 minutes with subsequent washing as above.
iv)
100 µl of substrate indicator system is added to all wells.
v)
The reaction is stopped after 15 minutes at room temperature by the addition of 25 µl of 37 mM NaCN.
Alternatively, commercially available detergents can be used after pretesting. The results are read
photometrically at a wavelength of 405 nm.
vi)
Calculation of results: absorbance (with antigen) minus absorbance (without antigen) = net extinction.
A reaction exceeding a net extinction of 0.3 is regarded as a positive result.
A competitive ELISA has also been described for detecting antibody against Trypanosoma equiperdum
(Katz et al., 2000).
d)
Other serological tests
Other serological tests have been used, including radioimmunoassay, counter immunoelectrophoresis and
agar gel immunodiffusion (AGID) tests (Caporale et al., 1981; Hagebock et al., 20199303). The AGID has
been used to confirm positive tests and to test anticomplementary sera. A seven-well pattern in 0.8%
agarose in Tris buffer is used, with the CF test antigen in the centre well and positive control sera and
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Chapter 2.5.3. — Dourine
unknown sera in alternate peripheral wells. A method has been published for diagnosing equine
piroplasmosis, glanders and dourine at the same time, using immunoblotting (Katz et al., 1999). A card
agglutination test has been developed that compares favourably with the CF test (Claes et al., 2005).
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
There are no biological products available. Control of the disease depends on compulsory notification and
slaughter of infected animals. Good hygiene at assisted matings is also essential.
REFERENCES
BARNER R.D. (1963). Protozoal diseases. In: Equine Medicine and Surgery, Bone J.F. et al., eds. American
Veterinary Publications, Santa Barbara, California, USA, 205–210.
BUNDESINSTITUT FÜR GESUNDHEITLICHEN VERBRAUCHERSCHUTZ UND VETERINÄRMEDIZIN (BgVV) (Federal Institute for
Consumer Health Protection and Veterinary Medicine) (1995). Working Protocols: ELISA on Dourine. BgVV. P.O.
Box 33 00 13, D-14191 Berlin, Germany.
CAPORALE V.P., BIANCIFIORI F., DI MATTEO A., NANNINI D. & URBANI G. (1981). Comparison of various tests for the
serological diagnosis of Trypanosoma equiperdum infection in the horse. Comp. Immunol. Microbiol. Infect. Dis.,
4, 243–246.
CLAES F., AGBO E.C., RADWANSKA M., TE PAS M.F., BALTZ T., DE WAAL D.T., GODDEERIS B.M., CLAASSEN E. &
BUSCHER P. (2003). How does T. equiperdum fit into the Trypanozooan genus? A cluster analysis and multiplex
genotyping approach. Parasitol., 126, 425–431.
CLAES F., ILGEKBAYEVA G.D., VERLOO D., SAIDOULDIN T.S., GEERTS S., BUSCHER P. & GODDEERIS B.M. (2005).
Comparison of serological tests for equine trypanosomosis in naturally infected horses from Kazakhstan. Vet.
Parasitol., 131 (3–4), 221–225.
HAGEBOCK J.M., CHIEVES L., FRERICHS W.M. & MILLER C.D. (1993). Evaluation of agar gel immunodiffusion and
indirect fluorescent antibody assays as supplemental tests for dourine in equids. Am. J. Vet. Res., 54, 1201–
1208.
HENNING M.W. (1955). Animal Diseases in South Africa, Third Edition. Central News Agency, South Africa.
HERR S., HUCHZERMAYER H.F.K.A., TE BRUGGE L.A., WILLIAMSON C.C., ROOS J.A. & SCHIELE G.J. (1985). The use of
a single complement fixation technique in bovine brucellosis, Johne’s disease, dourine, equine piroplasmosis and
Q fever serology. Onderstepoort J. Vet. Res., 52, 279–282.
HOARE C.A. (1972). The Trypanosomes of Mammals. A Zoological Monograph. Blackwell Scientific Publications,
Oxford & Edinburgh, UK.
KATZ J.B., CHIEVES L.P., HENNAGER S.G., NICHOLSON J.M., FISHER T.A. & BYERS P.E. (1999). Serodiagnosis of
equine piroplasmosis, dourine and glanders using an arrayed immunoblotting method. J. Vet. Diagn. Invest., 11,
292–294.
KATZ J.B., DEWALD R. & NICHOLSON J. (2000). Procedurally similar competitive immunoassay systems for the
serodiagnosis of Babesia equi, Babesia caballi, Trypanosoma equiperdum and Burkholderia mallei infection in
horses. J. Vet. Diagn. Invest., 12, 46–50.
LANHAM S.M. & GODFREY D.G. (1970). Isolation of salivarian trypanosomes from man and other mammals using
DEAE-cellulose. Exp. Parasitol., 28, 521–534.
MINISTRY OF AGRICULTURE, FISHERIES AND FOOD (1986). A Manual of Veterinary Parasitological Laboratory
Techniques, Third Edition. Her Majesty’s Stationery Office, London, UK.
UNITED STATES DEPARTMENT OF AGRICULTURE (2006). Complement Fixation Test for Detection of Antibodies to
Trypanosoma equiperdum – Microtitration Test. United States Department of Agriculture, Animal and Plant Health
Inspection Service, National Veterinary Services Laboratories, Ames, Iowa, USA.
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Chapter 2.5.3. — Dourine
WASSALL D.A., GREGORY R.J.F. & PHIPPS L.P. (1991). Comparative evaluation of enzyme-linked immunosorbent
assay (ELISA) for the serodiagnosis of dourine. Vet. Parasitol., 39, 233–239.
WATSON A.E. (1920). Dourine in Canada 1904–1920. History, Research and Suppression. Dominion of Canada
Department of Agriculture, Health of Animals Branch, Ottawa, Canada.
*
* *
NB: There is an OIE Reference Laboratory for Dourine
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests and reagents for dourine
OIE Terrestrial Manual 2012
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.4.
EPIZOOTIC LYMPHANGITIS
SUMMARY
Epizootic lymphangitis is a contagious, chronic disease of horses and other Equidae characterised
clinically by a spreading, suppurative, ulcerating pyogranulomatous dermatitis and lymphangitis.
This is seen particularly in the neck, legs and chest. It can also present as an ulcerating
conjunctivitis, or rarely a multifocal pneumonia. Transmission is by contact of infected material with
traumatised skin, by biting flies, ticks or inhalation. The causative agent, Histoplasma capsulatum
var. farciminosum, is a thermally dimorphic, fungal soil saprophyte. Differential diagnoses include
glanders (farcy), caused by Burkholderia mallei, ulcerative lymphangitis due to Corynebacterium
pseudotuberculosis, sporotrichosis caused by Sporothrix schenckii, and the skin lesions of
histoplasmosis caused by H. capsulatum var. capsulatum. Amphotericin B injection with local
wound drainage and inorganic iodides are used to treat early cases.
Identification of the agent: Identification of the agent is made by its appearance in smears of the
exudate or in histological sections of the lesion material. The yeast form of the organism is present
in large numbers in well established lesions, and appears as pleomorphic ovoid to globose
structures, approximately 2–5 µm in diameter, located both extracellularly and intracellularly in
macrophages and giant cells. Organisms are usually surrounded by a ‘halo’ when stained with
Gram stain, haematoxylin and eosin, Periodic acid–Schiff reaction or Gomori methenamine–silver
stain. The mycelial form of the organism grows slowly under aerobic conditions at 25–30°C on a
variety of media, including Mycobiotic agar, enriched Sabouraud’s dextrose agar, brain–heart
infusion agar, and pleuropneumonia-like organism nutrient agar. Conversion to the yeast phase at
37°C must be demonstrated.
Serological and other tests: Antibodies to H. capsulatum var. farciminosum develop at or before
the onset of clinical signs. Assays reported for detection of antibody include fluorescent antibody,
enzyme-linked immunosorbent assay, and passive haemagglutination tests. In addition, a skin
hypersensitivity test has been described.
Requirements for vaccines and diagnostic biologicals: Killed and live vaccines have been used
on a limited scale in endemic areas, but they are not readily available.
A. INTRODUCTION
Epizootic lymphangitis is a contagious, chronic disease of horses, mules and donkeys. The disease is
characterised clinically by a suppurative, ulcerating, and spreading pyogranulomatous, multifocal dermatitis and
lymphangitis. It is seen most commonly in the extremities, chest wall and the neck, but it can also be present as
an ulcerating conjunctivitis of the palpebral conjunctiva, or rarely as a multifocal pneumonia. The organism may
also invade open lesions including ruptured strangles abscesses and castration wounds. It has also been called
pseudofarcy or pseudoglanders. Another synonym is equine histoplasmosis, which may be a more accurate
name for the disease, as not all clinical cases present obvious lymphangitis. The form that the disease takes
seems to depend primarily on the route of entry (Singh, 1965). The traumatised skin is either infected directly by
infected pus, nasal or ocular excretions or indirectly by soil or contaminated harnesses, grooming equipment,
feeding and watering utensils, wound dressings or flies. It is also believed that ticks may play a role in the
transmission of this agent (Ameni & Terefe, 2004). The conjunctival form of the disease is believed to be spread
by flies of the Musca or Stomoxys genera (Singh, 1965). The pulmonary form of the disease is infrequent and is
presumed to occur after inhalation of the organism. The incubation period is from about 3 weeks to 2 months
(Ameni, 2006). In all cases, the lesions are nodular and granulomatous in character, and the organism, once
established, spreads locally by invasion and then via the lymphatics. There is often thickening, or ‘cording’, of
lymphatics, with the formation of pyogranulomatous nodules. Regional lymph nodes may by enlarged and
inflamed. Lesions usually heal spontaneously after 2–3 months, resulting in stellate scar formation. However,
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Chapter 2.5.4. — Epizootic lymphangitis
extensive lesions with high mortality rates can occur in areas where there is poor veterinary care and nutrition
(Ameni, 2006).
The causative agent, Histoplasma capsulatum var. farciminosum, is a thermally dimorphic fungus. The mycelial
form is present in soil; the yeast form is usually found in lesions. Histoplasma farciminosum was formerly
described as an independent species, but this assessment has been changed and it is now considered to be a
variety of H. capsulatum due to the close morphological similarities of both the mycelial and yeast forms (Ueda et
al, 2003). Antigenically, H. capsulatum var. farciminosum and H. capsulatum var. capsulatum are
indistinguishable, however the latter is the cause of disseminated histoplasmosis, is endemic in North America
and has a wide host range (Robinson & Maxie, 1993). DNA sequences of four protein-coding genes have been
analysed to elucidate the evolutionary relationships of H. capsulatum varieties. This indicated that H. capsulatum
var. farciminosum is deeply buried in the branch of SAm Hcc group A, (H60 to -64, -67, -71, -74 and -76), looking
as if it were an isolate of South American H. capsulatum var. capsulatum (Kasuga et al, 1999).
The cutaneous form of the disease may be confused with farcy (the skin form of glanders), which is caused by
Burkholderia mallei, ulcerative lymphangitis, which is caused by Corynebacterium pseudotuberculosis, indolent
ulcers acaused by Rhodococcus equi, sporotrichosis caused by Sporothrix schenckii, and histoplasmosis caused
by H. capsulatum var. capsulatum, cryptococcosis, strangles. sarcoids and cutaneous lymphosarcomas
(Jungerman & Schwartzman, 1972; Lehmann et al, 1996).
The disease is more common in the tropics and subtropics and is endemic in north, east and north-east Africa,
and some parts of Asia, including some countries bordering the Mediterranean Sea, India, Pakistan and Japan.
The disease is common in Ethiopia, especially in cart horses, affecting an average of 18.8% of horses in warm,
humid areas between 1500 and 2300 metres above sea level (Ameni, 2006; Ameni & Terefe, 2004). Reports from
other parts of the world are sporadic and all cases must be verified by laboratory testing. The prevalence of the
disease increases with assembling of animals; it was much more common, historically, when large numbers of
horses were stabled together for cavalry and other transportation needs. Mainly, it is horses, mules, and donkeys
that are affected by the disease, although infection may occur in camels, cattle and dogs (Ueda et al, 2003).
Experimentally, other animals are refractory to infection subsequent to inoculation, with the exception of certain
laboratory animal species such as mice, guinea-pigs and rabbits (Herve et al, 1994; Singh, 1965). Infection in
humans has also been reported (Al-Ani et al, 1998; Chandler et al, 1980; Guerin et al, 1992).
The disease is eradicated by the humane slaughter of infected horses, disinfection of infected premises and
restricting the movement of equids from infected premises. In endemic areas where eradication is not possible,
inorganic iodides can be used for therapy in early cases (Al-Ani, 1999). Localised nodules can also be lanced, the
pus drained and the nodules packed with a 7% tincture of iodine. If affordable, amphotericin B can be used.
As the clinical signs of epizootic lymphangitis can be confused with those of other diseases in the field, definitive
diagnosis rests on laboratory confirmation.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
Material should be collected directly from unruptured nodules. For microbiological isolation, the material should be
placed in a liquid nutrient medium with antibacterials and kept refrigerated until culturing, which should be
attempted as soon as possible. For direct examination, swabs of lesion material can be smeared on glass slides
and fixed immediately. For histopathology, sections of lesion material, including both viable and nonviable tissue,
should be placed in 10% neutral buffered formalin. Confirmation of the disease is dependent on the
demonstration of H. capsulatum var. farciminosum.
a)
Direct microscopic examination
•
Gram-stained smears
Smears can be stained directly with Gram’s stain and examined for the typical yeast form of the organism,
which will appear as Gram-positive, pleomorphic, ovoid to globose structures, approximately 2–5 µm in
diameter (Al-Ani et al., 1998). They may occur singly or in groups, and may be found either extracellularly or
within macrophages. A halo around the organisms (unstained capsule) is frequently observed.
•
Histopathology
In haematoxylin and eosin (H&E)-stained histological sections, the appearance of the lesion is quite
characteristic and consists of pyogranulomatous inflammation with fibroplasia. Langhans giant cells are
common. The presence of numerous organisms, both extracellularly and intracellularly within macrophages
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Chapter 2.5.4. — Epizootic lymphangitis
or multinucleated giant cells in tissue sections stained with H&E, Periodic acid–Schiff reaction and Gomori
methenamine–silver stain are observed (Robinson & Maxie, 1993). There is some indication that the number
of organisms increases with chronicity. The organisms are pleomorphic, often described as slightly lemonshaped basophilic masses, varying from 2 to 5 µm in diameter, that are surrounded by a ‘halo’ when stained
with H&E or Gram’s stain (Al-Ani, 1999).
•
Electron microscopy
Electron microscopy has been applied to skin biopsy samples of 1.5–2.0 mm immediately prefixed in
phosphate buffered 2% glutaraldehyde solution at 4°C and post-fixed in 1% osmium tetroxide. Ultra-thin
sections were cut and stained with uranyl acetate and lead citrate. Examination demonstrated the fine
internal structure of the organism, H. capsulatum var. farciminosum, including the cell envelope, plasma
membrane, cell wall, capsule and inner cell structures (Al-Ani, 1999).
b)
Culture
The mycelial form of H. capsulatum var. farciminosum grows slowly on laboratory media (2–8 weeks at
26°C). Media that can be used include Mycobiotic agar (Al-Ani et al, 1998), Sabouraud’s dextrose agar agar
enriched with 2.5% glycerol, brain–heart infusion agar supplemented with 10% horse blood, and
pleuropneumonia-like organism (PPLO) nutrient agar enriched with 2% dextrose and 2.5% glycerol, pH 7.8
(Guerin et al, 1992; Robinson & Maxie, 1993). The addition of antibiotics to the media is recommended:
cycloheximide (0.5 g/litre) and chloramphenicol (0.5 g/litre). Broad-spectrum antibacterial activity is obtained
if gentamicin (50 mg/litre) and penicillin G (6 × 106 units/litre) are used instead of chloramphenicol. Colonies
appear in 2–8 weeks as dry, grey-white, granular, wrinkled mycelia. The colonies become brown with aging.
Aerial forms occur, but are rare. The mycelial form produces a variety of conidia, including chlamydospores,
arthroconidia and some blastoconidia. However, the large round double-walled macroconidia that are often
observed in H. capsulatum var. capsulatum are lacking.
As a confirmatory test the yeast form of H. capsulatum var. farciminosum can be induced by subculturing
some of the mycelium into brain–heart infusion agar containing 5% horse blood or by using Pine’s medium
alone at 35–37°C in 5% CO2. Yeast colonies are flat, raised, wrinkled, white to greyish brown, and pasty in
consistency (Robinson & Maxie, 1993). However, complete conversion to the yeast phase may only be
achieved after four to five repeated serial transfers on to fresh media every 8 days.
c)
Animal inoculation
Experimental transmission of H. capsulatum var. farciminosum has been attempted in mice, guinea-pigs and
rabbits. Immunosuppressed mice are highly susceptible to experimental infection and can be used for
diagnostic purposes (Al-Ani, 1999).
2.
Serological tests
There are published reports of various tests to detect antibodies as well as a skin hypersensitivity test for
detection of cell-mediated immunity. Antibodies usually develop at or just after the onset of clinical signs.
a)
Fluorescent antibody tests
•
Indirect fluorescent antibody test
The following non-quantitative procedure is as described by Fawi (1969).
i)
Slides containing the organisms are made by smearing the lesion contents on to a glass slide or by
emulsifying the cultured yeast phase of the organism in a saline solution and creating a thin film on a
glass slide.
ii)
The slides are heat-fixed by passing the slide through a flame.
iii)
The slides are then washed in phosphate buffered saline (PBS) for 1 minute.
iv)
Undiluted test sera are placed on the slides, which are then incubated for 30 minutes at 37°C.
v)
The slides are washed in PBS three times for 10 minutes each.
vi)
Fluorescein isothiocyanate (FITC)-conjugated anti-horse antibody at an appropriate dilution is flooded
over the slides, which are then incubated for 30 minutes at 37°C.
vii)
Washing in PBS is repeated three times for 10 minutes each.
viii) The slides are examined using fluorescence microscopy.
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Chapter 2.5.4. — Epizootic lymphangitis
•
Direct fluorescent antibody test
The following procedure is as described by Gabal et al (1983).
b)
i)
The globulin fraction of the test serum is precipitated, and then re-suspended to its original serum
volume in saline. The serum is then conjugated to FITC.
ii)
Small colony particles of the cultured mycelial form of the organism are suspended in 1–2 drops of
saline on a glass slide. With a second slide, the colony particles are crushed and the solution is
dragged across the slide to create a thin film.
iii)
The smears are heat-fixed.
iv)
The slides are washed in PBS for 1 minute.
v)
The slides are incubated with dilutions of conjugated serum for 60 minutes at 37°C.
vi)
The slides are washed in PBS three times for 5 minutes each.
vii)
The slides are examined using fluorescence microscopy.
Indirect Enzyme-linked immunosorbent assay
The following procedure is as described by Gabal & Mohammed (1985).
i)
The mycelial form of the organism is produced on Sabouraud’s dextrose agar in tubes, and incubated
for 4 weeks at 26°C. Three colonies are ground in 50 ml of sterile PBS. The suspension is diluted
1/100 and the 96-well microtitre plates are coated with 100 µl/well.
ii)
The plates are incubated at 4°C overnight.
iii)
The plates are washed with PBS containing Tween 20 (0.5 ml/litre) (PBS-T) three times for 3 minutes
each.
iv)
The plates are incubated with 5% bovine serum albumin, 100 µl/well, at 23–25°C for 30 minutes, with
shaking.
v)
The plates are washed with PBS-T three times for 3 minutes each.
vi)
The sera are serially diluted using twofold dilution in duplicate in PBS-T, starting with a 1/50 dilution
and incubated for 30 minutes at 23–25°C.
vii)
The plates are washed with PBS-T three times for 3 minutes each.
viii) Peroxidase-labelled goat anti-horse IgG is diluted 1/800 and used at 100 µl/well, with incubation for
30 minutes at 23–25°C, with shaking.
c)
ix)
The plates are washed with PBS-T three times for 3 minutes each.
x)
Finally, 100 µl/well of hydrogen peroxide and ABTS (2,2’-Azino-di-[3-ethyl-benzthiazoline]-6-sulphonic
acid) in a citric acid buffer, pH 4, is added.
xi)
The plates are read at 60 minutes in a spectrophotometer at wavelength 405 nm.
xii)
The absorbance values are obtained twice from each serum dilution and the standard deviation and
average percentage of the absorbance values of the different serum samples are considered in the
interpretation of the results.
Passive haemagglutination test
The following procedure is as described by Gabal & Khalifa (1983).
i)
The organism is propagated for 8 weeks on Sabouraud’s dextrose agar. Five colonies are scraped,
ground, suspended in 200 ml of saline, and sonicated for 20 minutes. The remaining mycelial elements
are filtered out, and the filtrate is diluted 1/160.
ii)
Normal sheep red blood cells (RBCs) are washed, treated with tannic acid, washed, and re-suspended
as a 1% cell suspension.
iii)
Different dilutions of the antigen preparation are mixed with the tanned RBCs and incubated in a water
bath at 37°C for 1 hour. The RBCs are collected by centrifugation, washed three times in buffered
saline and re-suspended to make a 1% cell suspension.
iv)
Test sera are inactivated by heating at 56°C for 30 minutes and then absorbed with an equal volume of
washed RBCs.
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Chapter 2.5.4. — Epizootic lymphangitis
d)
v)
Dilutions of serum (0.5 ml) are placed in test tubes with 0.05 ml of antigen-coated tanned RBCs.
vi)
Agglutination is recorded at 2 and 12 hours.
vii)
Agglutination is detected when the RBCs form a uniform mat on the bottom of the tube. A negative test
is indicated by the formation of a ‘button’ of RBCs at the bottom of the tube.
Skin hypersensitivity tests
Two skin hypersensitivity tests for the diagnosis of epizootic lymphangitis have been described. The first test
was described by Gabal & Khalifa (1983) and adapted by Armeni et al (2006).
i)
A pure culture of H. farcimonsum is propagated for 8 weeks on Sabouraud’s dextrose agar containing
2.5% glycerol. Five colonies are scraped, ground, suspended in 200 ml of saline, undergo five freeze–
thaw cycles and are sonicated at an amplitude of 40° for 20 minutes. The remaining mycelial elements
are removed by centrifugation at 1006 g at 4°C for 11 minutes. Sterility of the preparation is verified by
incubating an aliquot on Sabouraud’s dextrose agar at 26°C for 4 weeks.
ii)
Animals are inoculated intradermally with 0.1 ml containing 0.2 mg/ml protein in the neck.
iii)
The inoculation site is examined for the presence of a local indurated and elevated area at 24–
48 hours post-injection. An increase in skin thickness of > 4 mm is considered to be positive.
Alternatively, a ‘histofarcin’ test has been described by Soliman et al (1985).
i)
The mycelial form of the organism is grown on polystyrene discs floating on 250 ml of PPLO media
containing 2% glucose and 2.5% glycerine at 23–25°C for 4 months.
ii)
The fungus-free culture filtrate is mixed with acetone (2/1) and held at 4°C for 48 hours.
iii)
The supernatant is decanted and the acetone is allowed to evaporate.
iv)
Precipitate is suspended to 1/10 original volume in distilled water.
v)
Animals are inoculated intradermally with 0.1 ml of antigen in the neck.
vi)
The inoculation site is examined for the presence of a local indurated and elevated area at 24, 48 and
72 hours post-injection.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
Control of the disease is usually through elimination of the infection. This is achieved by culling infected horses
and application of strict hygiene practices to prevent spread of the organism. There are published reports on the
use of killed (Al-Ani et al, 1998) and live attenuated vaccines (Zhang et al, 1986) in areas where epizootic
lymphangitis is endemic, apparently with relatively good results.
The antigens used for skin hypersensitivity testing are described in the previous section.
REFERENCES
AL-ANI F.K. (1999). Epizootic lymphangitis in horses: a review of the literature. Rev. sci. tech. Off. int. Epiz., 18,
691–699.
AL-ANI F.K., ALI A.H. & BANNA H.B. (1998). Histoplasma farciminosum infection of horses in Iraq. Veterinarski
Arhiv., 68, 101–107.
AMENI G. (2006). Preliminary trial on the reproducibility of epizootic lymphangitis through experimental infection of
two horses. Short Communication. Veterinary J., 172 (3), 553–555.
AMENI G. & TEREFE W. (2004). A cross-sectional study of epizootic lymphangitis in cart-mules in western Ethiopia.
Preventive Vet. Med., 66, 93–99.
AMENI G. TEREFE W. & HAILU A. (2006). Histofarcin test for the diagnosis of epizootic lymphanigitis in Ethiopia:
development, optimisation and validation in the field. Veterinary J., 171, 358–362.
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CHANDLER F.W., KAPLAN W. & AJELLO L. (1980). Histopathology of Mycotic Diseases. Year Book Medical
Publishers, Chicago, USA, 70–72 and 216–217.
FAWI M.T. (1969). Fluorescent antibody test for the serodiagnosis of Histoplasma farciminosum infections in
Equidae. Br. Vet. J., 125, 231–234.
GABAL M.A., BANA A.A. & GENDI M.E. (1983). The fluorescent antibody technique for diagnosis of equine
histoplasmosis (epizootic lymphangitis). Zentralbl. Veterinarmed. [B], 30, 283–287.
GABAL M.A. & KHALIFA K. (1983). Study on the immune response and serological diagnosis of equine
histoplasmosis (epizootic lymphangitis). Zentralbl. Veterinarmed. [B], 30, 317–321.
GABAL M.A. & MOHAMMED K.A. (1985). Use of enzyme-linked immunosorbent assay for the diagnosis of equine
Histoplasma farciminosi (epizootic lymphangitis). Mycopathologia, 91, 35–37.
GUERIN C., ABEBE S. & TOUATI F. (1992). Epizootic lymphangitis in horses in Ethiopia. J. Mycol. Med., 2, 1–5.
HERVE V., LE GALL-CAMPODONICO P., BLANC F., IMPROVISI, L., DUPONT, B, MATHIOT C. & LE GALL F. (1994).
Histoplasmose a Histoplasma farciminosum chez un cheval africain. J. Mycologie Med., 4, 54.
JUNGERMAN P.F. & SCHWARTZMAN R.M. (1972). Veterinary Medical Mycology. Lea & Febiger. Philadelphia, USA.
KASUGA T., TAYLOR T.W. & WHITE T.J. (1999). Phylogenetic relationships of varieties and geographical groups of
the human pathogenic fungus Histoplasma capsulatum darling. J. Clin. Microbiol., 37, 653–663.
LEHMANN P.F. HOWARD D.H. & MILLER J.D. (1996). Veterinary Mycology. Springer-Verlag, Berlin, Germany, 96,
251–263.
ROBINSON W.F. & MAXIE M.G. (1993). The cardiovascular system. In: Pathology of Domestic Animals, Vol. 3.
Academic Press, New York, USA, 82–84.
SINGH T. (1965). Studies on epizootic lymphangitis. I. Modes of infection and transmission of equine
histoplasmosis (epizootic lymphangitis). Indian J. Vet. Sci., 35, 102–110.
SOLIMAN R., SAAD M.A. & REFAI M. (1985). Studies on histoplasmosis farciminosii (epizootic lymphangitis) in Egypt.
III. Application of a skin test (‘histofarcin’) in the diagnosis of epizootic lymphangitis in horses. Mykosen, 28, 457–
461.
UEDA Y., SANO A. TAMURA M., INOMATA T., KAMEI K., YOKOYAMA K., KISHI F., ITO J., Y., MIYAJI M. & NISHIMURA K.
(2003). Diagnosis of histoplasmosis by detection of the internal transcribed spacer region of fungal rRNA gene
from a paraffin-embedded skin sample from a dog in Japan. Vet. Microbiol., 94, 219–224.
ZHANG W.T., WANG Z.R., LIU Y.P., ZHANG D.L., LIANG P.Q., FANG Y.Z., HUANG Y.J. & GAO S.D. (1986). Attenuated
vaccine against epizootic lymphangitis in horses. Chinese J. Vet. Sci. Tech., 7, 3–5.
*
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CHAPTER 2.5.5.
EQUINE ENCEPHALOMYELITIS
(Eastern and Western)
SUMMARY
Eastern and Western equine encephalomyelitis viruses belong to the genus Alphavirus of the family
Togaviridae. Alternate infection of birds and mosquitoes maintain these viruses in nature. The
disease occurs sporadically in horses and humans from mid-summer to late autumn. Horses and
humans are tangential dead-end hosts. The disease in horses is characterised by fever, anorexia,
and severe depression. In severe cases, the disease in horses progresses to hyperexcitability,
blindness, ataxia, severe mental depression, recumbency, convulsions, and death. Eastern equine
encephalomyelitis (EEE) virus infection in horses is often fatal, while Western equine
encephalomyelitis (WEE) virus can cause a subclinical or mild disease with less than 30% mortality.
EEE and WEE have been reported to cause disease in poultry, game birds and ratites. Sporadic
cases of EEE have been reported in cows, sheep, pigs, deer, and dogs.
Identification of the agent: A presumptive diagnosis of EEE or WEE can be made when
susceptible horses display the characteristic somnolence and other signs of neurological disease in
areas where haematophagous insects are active. There are no characteristic gross lesions.
Histopathological lesions can provide a presumptive diagnosis. EEE virus can usually be isolated
from the brain and sometimes other tissues of dead horses, however WEE virus is rarely isolated.
EEE and WEE viruses can be isolated from field specimens by inoculating newborn mice,
embryonating chicken eggs, cell cultures, or newly hatched chickens. The virus is identified by
complement fixation (CF), immunofluorescence, or plaque reduction neutralisation (PRN) tests.
EEE and WEE viral RNA may also be detected by reverse-transcription polymerase chain reaction
methods.
Serological tests: Antibody can be identified by PRN, haemagglutination inhibition (HI), CF tests,
or IgM capture enzyme-linked immunosorbent assay.
Requirements for vaccines and diagnostic biologicals: EEE and WEE vaccines are safe and
immunogenic. They are produced in cell culture and inactivated with formalin.
A. INTRODUCTION
Eastern equine encephalomyelitis (EEE) and Western equine encephalomyelitis (WEE) viruses are members of
the genus Alphavirus of the family Togaviridae. The natural ecology for virus maintenance occurs via alternate
infection of birds and ornithophilic mosquitoes. Clinical disease may be observed in humans and horses, both of
which are dead-end hosts for these agents. EEE has been diagnosed in Quebec and Ontario in Canada, central
and eastern regions of the United States of America (USA), the Caribbean Islands, Mexico, and Central and
South America. Disease caused by the WEE virus has been reported in the western USA and Canada, Mexico,
and Central and South America (Morris, 1989; Reisen & Monath, 1989; Walton et al., 1981). Highlands J virus,
antigenically related to WEE virus, has been isolated in eastern USA. Although Highlands J virus is generally
believed not to cause disease in mammals, it has been isolated from the brain of a horse dying of encephalitis in
Florida (Karabatsos et al., 1988).
Even though the mortality is lower for WEE, the clinical signs of EEE and WEE can be identical. The disease
caused by either virus is also known as sleeping sickness. Following an incubation period of 5–14 days, clinical
signs include fever, anorexia, and depression. A presumptive diagnosis of EEE or WEE virus infection in
unvaccinated horses can be made if the characteristic somnolence is observed during the summer in temperate
climates or the wet season in tropical and subtropical climates, when the mosquito vector is plentiful. However, a
number of other diseases, such as West Nile virus and Venezuelan equine encephalomyelitis (chapters 2.1.20
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Chapter 2.5.5. — Equine encephalomyelitis (Eastern and Western)
and 2.5.14, respectively), produce similar clinical signs and the diagnosis must be confirmed by the described
diagnostic test methods. WEE virus infection in horses is often observed over a wide geographical area, e.g.
sporadic cases over 1000 square miles. EEE virus infections are usually observed in limited geographical areas.
Isolated events of high mortality in captive-raised game birds, primarily pheasants, chukars, aquarium penguins,
and quail have been traced to WEE, EEE, or Highlands J virus infection (Morris, 1989; Reisen & Monath, 1989;
Tuttle et al., 2005). Most encephalomyelitis infections in domestic fowl are caused by EEE virus and occur on the
east coast states of the USA. The virus is introduced by mosquitoes, but transmission within the flocks is primarily
by feather picking and cannibalism. Both EEE and WEE viruses have caused a fatal disease in ratites.
Haemorrhagic enteritis has been observed in emus infected with EEE and WEE viruses, and morbidity and
mortality rates may be greater than 85%. Highlands J and EEE viruses have been found to produce depression,
somnolence, decreased egg production, and increased mortality in turkeys (Guy, 1997). EEE virus has been
reported to cause disease in cows (McGee et al., 1992; Pursell et al., 1976), sheep (Bauer et al., 2005), pigs
(Elvinger et al., 1996), white-tailed deer (Tate et al., 2005), and dogs (Farrar et al., 2005).
EEE virus causes severe disease in humans with a mortality rate of 30–70% and a high frequency of permanent
sequelae in patients who survive. WEE is usually mild in adult humans, but can be a severe disease in children.
The fatality rate is between 3 and 14%. Severe infection and death caused by EEE and WEE viruses have been
reported in laboratory workers; therefore, any work with these viruses must be performed at containment level 3
(see Chapter 1.1.3 Biosafety and biosecurity in the veterinary microbiology laboratory and animal facilities). It is
recommended that personnel be immunised against EEE and WEE viruses (United States Department of Health
and Human Services, 1999). Precautions should also be taken to prevent human infection when performing postmortem examinations on horses suspected of being infected with the equine encephalomyelitis viruses.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
The most definitive method for diagnosis of EEE or WEE is the isolation of the viruses. EEE virus can usually be
isolated from the brains of horses, unless more than 5 days have elapsed between the appearance of clinical
signs and the death of the horse. EEE virus can frequently be isolated from brain tissue even in the presence of a
high serum antibody titre. WEE virus is rarely isolated from tissues of infected horses. Brain is the tissue of choice
for virus isolation, but the virus has been isolated from other tissues, such as the liver and spleen. It is
recommended that a complete set of these tissues be collected in duplicate, one set for virus isolation and the
other set in formalin for histopathological examination. Specimens for virus isolation should be sent refrigerated if
they can be received in the laboratory within 48 hours of collection; otherwise, they should be frozen and sent with
dry ice. A complete set of tissues will allow the performance of diagnostic techniques for other diseases. For
isolation, a 10% suspension of tissue is prepared in phosphate buffered saline (PBS), pH 7.8, containing bovine
serum albumin (BSA) (fraction V; 0.75%), penicillin (100 units/ml), and streptomycin (100 µg/ml). The suspension
is clarified by centrifugation at 1500 g for 30 minutes.
EEE and WEE viruses can be isolated in a number of cell culture systems. The most commonly used cell cultures
are primary chicken or duck embryo fibroblasts, continuous cell lines of African green monkey kidney (Vero),
rabbit kidney (RK-13), or baby hamster kidney (BHK-21). Isolation is usually attempted in 25 cm2 cell culture
flasks. Confluent cells are inoculated with 1.0 ml of tissue suspension. Following a 1–2-hour absorption period,
maintenance medium is added. Cultures are incubated for 6–8 days, and one blind passage is made. EEE and
WEE viruses will produce a cytopathic change in cell culture. Cultures that appear to be infected are frozen. The
fluid from the thawed cultures is used for virus identification.
The newborn mouse is also considered to be a sensitive host system. Inoculate intracranially one or two litters of
1–4-day-old mice with 0.02 ml of inoculum using a 26-gauge 3/8 inch (9.3 mm) needle attached to a 1 ml
tuberculin syringe. The inoculation site is just lateral to the midline into the midportion of one lateral hemisphere.
Mice are observed for 10 days. Mice that die within 24 hours of inoculation are discarded. From 2 to 10 days
postinoculation, dead mice are collected daily and frozen at –70°C. Mouse brains are harvested for virus
identification by aspiration using a 20-gauge 1 inch (2.5 cm) needle attached to a 1 ml tuberculin syringe. A
second passage is made only if virus cannot be identified from mice that die following inoculation.
The chicken embryo is considered to be less sensitive than newborn mice when used for primary isolation of EEE
and WEE viruses. Tissue suspensions can be inoculated by the yolk-sac route into 6–8-day-old embryonating
chicken eggs. There are no diagnostic signs or lesions in the embryos infected with these viruses. Inoculated
embryos should be incubated for 7 days, but deaths usually occur between 2 and 4 days post-inoculation. Usually
only one passage is made unless there are dead embryos from which virus cannot be isolated. Newly hatched
chickens are susceptible and have been used for virus isolation. If this method is used, precautions must be taken
to prevent aerosol exposure of laboratory personnel, as infected birds can shed highly infectious virus.
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Chapter 2.5.5. — Equine encephalomyelitis (Eastern and Western)
EEE or WEE viruses can be identified in infected mouse or chicken brains, cell culture fluid, or amnionic-allantoic
fluid by complement fixation. A 10% brain suspension is prepared in veronal (barbitone) buffer; egg and cell
culture fluids are used undiluted or diluted 1/10 in veronal buffer. The fluid or suspension is centrifuged at 9000 g
for 30 minutes, and the supernatant fluid is tested against hyperimmune serum or mouse ascitic fluid prepared
against EEE and WEE viruses using a standard CF procedure (United States Department Of Health, Education
and Welfare, 1974). The CF test requires the overnight incubation at 4°C of serum-antigen with 7 units of
complement. Virus can be identified in cell culture by direct immunofluorescent staining. The less commonly used
method of virus identification is the neutralisation test, as outlined below.
Reverse-transcription polymerase chain reaction (RT-PCR) methods to detect EEE, WEE and VEE viral nucleic
acid in mosquitoes and vertebrate tissues have been described, although few have been extensively validated for
mammalian samples (Lambert et al., 2003; Linssen et al., 2000; Monroy et al., 1996; Vodkin et al., 1993). A
multiplex PCR method was developed to expedite differential diagnosis in cases of suspected EEE or West Nile
arboviral encephalomyelitis in horses (Johnson et al., 2003). The assay has enhanced speed and sensitivity
compared with cell culture virus isolation and has been used effectively in the USA during several recent
arbovirus seasons. Recently, a combination of an RT-PCR with an enzyme-linked immunosorbent assay (ELISA:
RT-PCR-ELISA) was reported as a method to identify alpha-viruses that are pathogenic to humans (Wang et al.,
2006).
Antigen-capture ELISA has been developed for EEE surveillance in mosquitoes. This can be used in countries
that do not have facilities for virus isolation or PCR (Brown et al., 2001). Immunohistochemical procedures for
diagnosis of EEE have also been described (Patterson et al., 1996).
2.
Serological tests
Serological confirmation of EEE or WEE virus infection requires a four-fold or greater increase or decrease in
antibody titre in paired serum samples collected 10–14 days apart. Most horses infected with EEE or WEE virus
have a high antibody titre when clinical disease is observed. Consequently, a presumptive diagnosis can be made
if an unvaccinated horse with appropriate clinical signs has antibody against only EEE or WEE virus. The
detection of IgM antibody by the ELISA can also provide a presumptive diagnosis of acute infection (Sahu et al.,
1994). The plaque reduction neutralisation (PRN) test or, preferably, a combination of PRN and
haemagglutination inhibition (HI) tests is the procedure most commonly used for the detection of antibody against
EEE and WEE viruses. There may be cross-reactions between antibody against EEE and WEE virus in the CF
and HI tests. CF antibody against both EEE and WEE viruses appears later and does not persist; consequently, it
is less useful for the serological diagnosis of disease.
a)
Complement fixation
The CF test is frequently used for the demonstration of antibodies, although the antibodies detected by the
CF test may not persist for as long as those detected by the HI or PRN tests. A sucrose/acetone mouse
brain extract is commonly used as antigen. The positive antigen is inactivated by treatment with 0.1% betapropiolactone.
In the absence of an international standard serum, the antigen should be titrated against a locally prepared
positive control serum. The normal antigen, or control antigen, is mouse brain from uninoculated mice
similarly extracted and diluted.
Sera are diluted 1/4 in veronal buffered saline containing 1% gelatin (VBSG), and inactivated at 56°C for
30 minutes. Titrations of positive sera may be performed using additional twofold dilutions. The CF antigens
and control antigen (normal mouse brain) are diluted in VBSG to their optimal amount of fixation as
determined by titration against the positive sera; guinea-pig complement is diluted in VBSG to contain
5 complement haemolytic units-50% (CH50). Sera, antigen, and complement are reacted in 96-well roundbottom microtitre plates at 4°C for 18 hours. The sheep red blood cells (SRBCs) are standardised to 2.8%
concentration. Haemolysin is titrated to determine the optimal dilution for the lot of complement used.
Haemolysin is used to sensitise 2.8% SRBCs and the sensitised cells are added to all wells on the microtitre
plate. The test is incubated for 30 minutes at 37°C. The plates are then centrifuged (200 g), and the wells
are scored for the presence of haemolysis. The following controls are used: (a) serum and control serum
each with 5 CH50 and 2.5 CH50 of complement; (b) CF antigen and control antigen each with 5 CH50, and
2.5 CH50 of complement; (c) complement dilutions of 5 CH50, 2.5 CH50, and 1.25 CH50; and (d) cell control
wells with only SRBCs and VBSG diluent. These controls test for anticomplementary serum and
anticomplementary antigen, activity of complement used in the test, and integrity of the SRBC indicator
system in the absence of complement, respectively.
To avoid anticomplementary effects, sera should be separated from the blood as soon as possible. Positive
and negative control sera should be used in the test.
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b)
Haemagglutination inhibition
The antigen for the HI test is the same as described above for the CF test. The antigen is diluted so that the
amount used in each haemagglutinating unit (HAU) is from four to eight times that which agglutinates 50% of
the RBCs in the test system. The haemagglutination titre and optimum pH for each antigen are determined
with goose RBCs diluted in pH solutions ranging from pH 5.8 to pH 6.6, at 0.2 intervals.
Sera are diluted 1/10 in borate saline, pH 9.0, and then inactivated at 56°C for 30 minutes. Kaolin treatment is
used to remove nonspecific serum inhibitors. Alternatively, nonspecific inhibitors may be removed by acetone
treatment of serum diluted 1/10 in PBS followed by reconstitution in borate saline. Sera should be absorbed
before use by incubation with a 0.05 ml volume of washed packed goose RBCs for 20 minutes at 4°C.
Following heat inactivation, kaolin treatment and absorption, twofold dilutions of the treated serum are
prepared in borate saline, pH 9.0 with 0.4% bovalbumin. Serum dilutions (0.025 ml/well) are prepared in a
96-well round-bottom microtitre plate in twofold dilutions in borate saline, pH 9.0, with 0.4% bovalbumin.
Antigen (0.025 ml/well) is added to the serum. Plates are incubated at 4°C overnight. RBCs are derived from
normal white male geese1 and washed three times in dextrose/gelatin/veronal (DGV), and a 7.0%
suspension is prepared in DGV. The 7.0% suspension is then diluted 1/24 in the appropriate pH solution,
and 0.05 ml per well is added immediately to the plates. Plates are incubated for 30 minutes at 37°C.
Positive and negative control sera are incorporated into each test. A test is considered to be valid only if the
control sera give the expected results. Titres of 1/10 and 1/20 are suspect, and titres of 1/40 and above are
positive.
c)
Enzyme-linked immunosorbent assay
The ELISA is performed by coating flat-bottomed plates with anti-equine IgM capture antibody (Sahu et al.,
1994). The antibody is diluted according to the manufacture’s recommendations in 0.5 M carbonate buffer,
pH 9.6, and 50 µl is added to each well. The plates are incubated at 37°C for 1 hour, and then at 4°C
overnight. Prior to use, the coated plates are washed three times with 200–300 µl/well of 0.01 M PBS
containing 0.05% Tween 20. After the second wash, 200 µl/well of PBS/Tween/5% nonfat dried milk is
added and the plates are incubated at room temperature for 1 hour. Following incubation, the plates are
washed again three times with PBS/Tween. Test and control sera are diluted 1/400 in 0.01 M PBS, pH 7.2,
containing 0.05% Tween 20, and 50 µl is added to each well. The plates are incubated at 37°C for
90 minutes and then washed three times. Next, 50 µl of viral antigen is added to all wells. (The dilution of the
antigen will depend on the source and should be empirically determined.) The plates are incubated overnight
at 4°C, and washed three times. Then, 50 µl of horseradish-peroxidase-conjugated monoclonal antibody
(MAb) to encephalitis virus2 is added. The plates are incubated for 90 minutes at 37°C and then washed six
times. Finally, 50 µl of freshly prepared ABTS (2,2’-Azino-bis-[3-ethylbenzo-thiazoline-6-sulphonic acid])
substrate + hydrogen peroxidase is added, and the plates are incubated at room temperature for 15–
40 minutes The absorbance of the test serum is measured at 405 nm. A test sample is considered to be
positive if the absorbance of the test sample in wells containing virus antigen is at least twice the
absorbance of negative control serum in wells containing virus antigen and at least twice the absorbance of
the sample tested in parallel in wells containing normal antigen.
d)
Plaque reduction neutralisation
The PRN test is very specific and can be used to differentiate between EEE and WEE virus infections. The
PRN test is performed in duck embryo fibroblast, Vero, or BHK-21 cell cultures in 25 cm2 flasks or six-well
plates. Volumes listed are for flasks; the volume should be halved for wells in six-well plates. The sera can
be screened at a 1/10 and 1/100 final dilution. Endpoints can be established using the PRN or HI test.
Serum used in the PRN assay is tested against 100 plaque-forming units (PFU) of virus (50 PFU for six-well
plates). The virus/serum mixture is incubated at 37°C for 75 minutes before inoculation on to confluent cell
culture monolayers in 25 cm2 flasks. The inoculum is adsorbed for 1 hour, followed by the addition of 6 ml of
overlay medium. The overlay medium consists of two solutions that are prepared separately. Solution I
contains 2 × Earle’s Basic Salts Solution without phenol red, 4% fetal bovine serum, 100 µg/ml gentamicin,
200 µg/ml nystatin, 0.45% solution of sodium bicarbonate, and 0.002% neutral red. When duck embryo
fibroblasts are used, Solution 1 also contains 6.6% yeast extract lactalbumin hydrolysate. Solution II consists
of 2% Noble agar that is sterilised and maintained at 47°C. Equal volumes of solutions I and II are adjusted
to 47°C and mixed together just before use. The test is incubated for 48–72 hours, and endpoints are based
on a 90% reduction in the number of plaques compared with the virus control flasks, which should have
about 100 plaques.
1
2
RBCs from adult domestic white male geese are preferred, but RBCs from other male geese can be used. If cells from
female geese are used, there may be more test variability. It has been reported that rooster RBCs cause a decrease in
the sensitivity of the test.
Available from: Centers for Disease Control and Prevention, Biological Reference Reagents, 1600 Clifton Road NE, Mail
Stop C21, Atlanta, Georgia 30333, United States of America.
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C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
Inactivated vaccines against EEE and WEE viruses are available commercially. Attenuated EEE and WEE virus
vaccines have not proven satisfactory. The vaccines licensed for use in the USA are prepared using the following
combinations: EEE and WEE; EEE, WEE, and Venezuelan equine encephalomyelitis (VEE); and EEE and VEE.
In addition, tetanus toxoid, inactivated influenza virus, and inactivated West Nile virus have been combined with
EEE and WEE or EEE, WEE, and VEE. Early vaccines were produced from virus propagated in embryonating
chicken eggs and inactivated with formalin. Current vaccines are prepared from virus propagated in cell culture,
and inactivated with formalin (Maire et al., 1970) or monoethylamine.
1.
Seed management
a)
Characteristics of the seed
Standard strains of EEE and WEE viruses that were isolated over 20 years ago have been used for vaccine
production and have been proven to produce a protective immunity. Strains of EEE virus that differ
antigenically and in molecular structure have been identified from different geographical regions. However,
the North American and Caribbean isolates appear to be similar (Weaver et al., 1994). Strains of WEE virus
isolated from different countries have been found to be similar both by MAb testing and RNA oligonucleotide
fingerprinting analysis (Reisen & Monath, 1989). A recent well-characterised isolate from the country where
the vaccine is to be used would be advantageous. Viruses that are selected must be immunogenic and
replicate to high titres in cell culture.
b)
Method of culture
Primary chicken embryo fibroblasts and Vero cells have been used for propagation of viruses used for
vaccine production. The fibroblasts should be prepared from specific pathogen free embryos. Other
susceptible cell lines could also be used.
c)
Validation as a vaccine
If a cell line is used, the master cell stock is tested to confirm the identity of the cell line, species of origin,
and freedom from extraneous agents. If primary cell cultures are used, a monolayer from each batch of each
subculture should be tested for extraneous agents including bacteria, fungi, mycoplasma, and viruses. The
master seed virus should also be tested to ensure freedom from bacteria, fungi, mycoplasma, and
extraneous viruses (see Chapter 1.1.7 Tests for sterility and freedom from contamination of biological
materials for a description of the method).
The vaccines are administered by the intramuscular (in most cases) or intradermal route in the cervical
region in two doses given 2–4 weeks apart. Annual revaccination is recommended. All foals vaccinated
before 1 year of age should be revaccinated before the next vector season.
2.
Method of manufacture
Details of the manufacture of vaccines currently on the market are not available. Consequently, the information
provided here is intended only as background reference material on the vaccines and not as a method of
manufacture. The virus and cell culture system should be selected so that a high virus titre, ≥106 TCID50 (50%
tissue culture infective dose) per ml, is obtained in under 48 hours. Virus for vaccine production can be prepared
from the supernatant fluid from infected cell cultures. The fluid is harvested when 70–100% of the monolayers
have the characteristic cytopathic changes. The virus titre is determined by titration in cell culture or mice. The
fluid is clarified by low speed centrifugation and filtered through gauze. The virus is inactivated by adding formalin
to a final concentration of 1/2000 (0.05%) and holding at 37°C for 24 hours. Residual formaldehyde is neutralised
by sodium bisulphite (Maire et al., 1970). The residual free formalin content in the inactivated vaccine should not
exceed 0.2% formaldehyde.
3.
In-process control
Cultures should be examined daily for virus-induced cytopathic effect. The harvested virus should be tested for
microbial contamination. The efficacy of the inactivation process should be checked by testing for viable virus.
4.
Batch control
a)
Sterility
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
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b)
Safety
The inactivated vaccine is safety tested by inoculating subcutaneously at least ten 6- to 12-hour-old chickens
with 0.5 ml of the vaccine. The chickens are observed each day for 10 days for unfavourable reactions that
are attributable to the vaccine (United States Code of Federal Regulations, 2000). Safety testing can also be
carried out by inoculating intracerebrally at least eight 1–4-day-old mice with 0.02 ml of the vaccine, and
observing for 7 days. It is critical that safety tests be conducted on each lot of vaccine to insure that there is
no residual virulent virus present.
c)
Potency
Potency testing is performed by inoculating each of ten guinea pigs with either EEE or WEE virus, using
one-half the horse dose on two occasions, 14–21 days apart, by the route recommended for the horse.
Serum samples from each vaccinate and each control are tested 14–21 days after the second dose using
the PRN test. The EEE titres should be ≥1/40, and the WEE titres should be ≥1/40 (US Code of Federal
Regulations), using Vero cells. If duck embryo fibroblasts are used in the PRN test, the titres will be lower.
An alternative potency test is to use intracerebral challenge, 14–21 days after the second vaccination. Each
guinea pig is inoculated with 0.1 ml of virus containing 100 LD50 (50% lethal dose). Simultaneous titration is
carried out. In order for the vaccine to be approved, 80% of the guinea pigs must survive both viruses.
d)
Duration of immunity
Comprehensive studies on duration of immunity are not available. An annual revaccination is recommended.
Foals that are vaccinated before 1 year of age should be revaccinated before the next vector season.
e)
Stability
The lyophilised vaccine is stable and immunogenic for 3 years if kept refrigerated at 2–7°C. After 3 years,
vaccine should be discarded. The vaccines should be used immediately after reconstitution.
f)
Preservatives
The preservatives used are thimerosal at a 1/10,000 dilution and antibiotics (neomycin, polymyxin
amphotercin B and gentamicin).
g)
Precautions (hazards)
Severe infection and death caused by EEE and WEE viruses have been reported in laboratory workers;
therefore, any work with these viruses must be carried out at least in a containment level 3 laboratory (see
chapter 1.1.3) using biological safety cabinets, and it is recommended that personnel be immunised against
EEE and WEE viruses (United States Department of Health and Human Services, 1999).
Pregnant mares and foals under 2 weeks old should not be vaccinated.
5.
Tests on the final product
a)
Safety
See Section C.4.b.
b)
Potency
See Section C.4.c.
REFERENCES
BAUER R.W., GILL M.S., POSTON R.B. & KIM D. Y. (2005). Naturally occurring eastern equine encephalitis in a
Hampshire wether. J. Vet. Diagn. Invest., 17, 281–285.
BROWN T.M., MITCHELL C.J., NASCI R.S., SMITH G.C. & ROEHRIG J.T. (2001). Detection of eastern equine
encephalitis virus in infected mosquitoes using a monoclonal antibody-based antigen-capture enzyme-linked
immunosorbent assay. Am. J. Trop. Med. Hyg., 65, 208–213.
ELVINGER F., BALDWIN C.A., LIGGETT A.D., TANK K.N., & STALLKNECHT D.E. (1996). Prevalence of exposure to
eastern equine encephalomyelitis virus in domestic and feral swine in Georgia. J. Vet. Diagn. Invest., 8, 481–484.
FARRAR M.D., MILLER D. L., BALDWIN C. A., STIVER S. L. & HALL C. L. (2005). Eastern equine encephalitis in dogs. J.
Vet. Diagn. Invest., 17, 614–617.
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GUY J.S. (1997). Arbovirus Infections. In: Diseases of Poultry, Calnek B.W., Barnes H.J., Beard C.W., McDougald
L.R., & Saif Y.M., ed. Iowa State University Press, Ames, Iowa, USA, 765–772.
JOHNSON D.J., OSTLUND E.N. & SCHMITT B.J. (2003). Nested multiplex RT-PCR for detection and differentiation of
West Nile virus and eastern equine encephalomyelitis virus in brain tissues. J. Vet. Diagn. Invest., 15, 488–493.
KARABATSOS N., LEWIS A.L., CALISHER C.H., HUNT A.R. & ROEHRIG J.T. (1988). Identification of Highland J virus from
a Florida horse. Am. J. Trop. Med. Hyg., 39, 603–606.
LAMBERT A.J., MARTIN D.A. & LANCIOTTI R.S. (2003). Detection of North American eastern and western equine
encephalitis viruses by nucleic acid amplification assays. J. Clin. Microbiol., 41, 379–385.
LINSSEN B., KINNEY R.M., AGUILAR P., RUSSELL K.L., WATTS D.M., KAADEN O.R. & PFEFFER M. (2000). Development
of reverse transcription-PCR assays specific for detection of equine encephalitis viruses. J. Clin. Microbiol., 38,
527–535.
MAIRE L.F. III, MCKINNEY R.W. & COLE F.E. Jr (1970). An inactivated eastern equine encephalomyelitis vaccine
propagated in chick-embryo cell culture. I. Production and testing. Am. J. Trop. Med. Hyg., 19, 119–122.
MCGEE E.D., LITTLETON C.H., MAPP J.B. & BROWN R.J. (1992). Eastern equine encephalomyelitis in an adult cow.
Vet. Pathol., 29, 361–363.
MONROY A.M., SCOTT T.W. & WEBB B.A. (1996). Evaluation of reverse transcriptase polymerase chain reaction for
the detection of eastern equine encephalomyelitis virus during vector surveillance. J. Med. Entomol., 33, 449–
457.
MORRIS C.D. (1989). Eastern equine encephalomyelitis. In: The Arboviruses: Epidemiology and Ecology, Vol. 3,
Monath T.P., ed. CRC Press, Boca Raton, Florida, USA, 1–12.
PURSELL A.R., MITCHELL F.E. & SEIBOLD H.R. (1976). Naturally occurring and experimentally induced eastern
encephalomyelitis in calves. J. Am. Vet. Med. Assoc., 169, 1101–1103.
PATTERSON J.S., MAES R.K., MULLANEY T.P., & BENSON C.L. (1996). Immunohistochemical Diagnosis of Eastern
Equine Encephalomyelitis. J. Vet. Diagn. Invest., 8, 156–160.
REISEN W.K. & MONATH T.P. (1989). Western equine encephalomyelitis. In: The Arboviruses: Epidemiology and
Ecology, Vol. 5, Monath T.P., ed. CRC Press, Boca Raton, Florida, USA, 89–137.
SAHU S.P., ALSTAD A.D., PEDERSEN D.D. & PEARSON J.E. (1994). Diagnosis of eastern equine encephalomyelitis
virus infection in horses by immunoglobulin M and G capture enzyme-linked immunosorbent assay. J. Vet. Diagn.
Invest., 6, 34–38.
TATE C.M., HOWERTH E.W., STALLKNECHT D.E., ALLISTION, A.B., FISHER J.R. & MEAD D.G. (2005). Eastern equine
encephalitis in a free-ranging white-tailed deer (Odocoileus virginianus). J. Wildl. Dis., 41, 241–245.
TUTTLE A.D., ANDREADIS T.G., FRASCA S. JR & DUNN J.L. (2005). Eastern equine encephalitis in a flock of African
penguins maintained at an aquarium. J. Am. Vet. Med. Assoc., 226, 2059–2062.
UNITED STATES CODE OF FEDERAL REGULATIONS (2000). Encephalomyelitis vaccine: Eastern and Western killed
virus. Title 9, Part 113, Section 113.207. US Government Printing Office, Washington DC, USA, 601–602.
UNITED STATES DEPARTMENT OF HEALTH, EDUCATION AND WELFARE (1974). A Guide to the Performance of the
Standardised Complement Fixation Method and Adaptation to Micro Test. Centers for Disease Control, Atlanta,
Georgia, USA.
UNITED STATES DEPARTMENT OF HEALTH AND HUMAN SERVICES (1999). Biosafety in Microbiological and Biomedical
Laboratories. US Government Printing Office, Washington DC, USA, 183–184.
VODKIN M.H., MCLAUGHLIN G.L., DAY J.F., SHOPE R.E. & NOVAK R.J. (1993). A rapid diagnostic assay for eastern
equine encephalomyelitis viral-RNA. Am. J. Trop. Med. Hyg., 49, 772–776.
WALTON T.E. (1981). Venezuelan, eastern, and western encephalomyelitis. In: Virus Diseases of Food Animals. A
World Geography of Epidemiology and Control. Disease Monographs, Vol. 2, Gibbs E.P.J., ed. Academic Press,
New York, USA, 587–625.
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WANG E., PAESSLER S., AGUILAR P.V., CARRARA A.S., NI H., GREENE I.P. & WEAVER S.C. (2006). Reverse
transcription-PCR-enzyme-linked immunosorbent assay for rapid detection and differentiation of alphavirus
infections. J. Clin. Microbiol., 44, 4000–4008.
WEAVER S.C., HAGENBAUGH A., BELLEW L.A., GOUSSET L., MALLAMPALLI V., HOLLAND J.J. & SCOTT T.W. (1994).
Evolution of Alphaviruses in the eastern equine encephalomyelitis complex. J. Virol., 68, 158–169.
*
* *
NB: There is an OIE Reference Laboratory for Equine encephalomyelitis (Eastern and Western):
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for equine encephalomyelitis (Eastern and Western)
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.6.
EQUINE INFECTIOUS ANAEMIA
SUMMARY
Equine infectious anaemia (EIA) is a persistent viral infection of equids. The causative agent, EIA
virus (EIAV) is a lentivirus in the family Retroviridae, subfamily Orthoretrovirinae. Other members of
the lentivirus genus include: bovine immunodeficiency virus; caprine arthritis encephalitis virus;
feline immunodeficiency virus; human immunodeficiency virus 1; human immunodeficiency virus 2;
and maedi/visna virus. EIA can be diagnosed on the basis of clinical signs, pathological lesions,
serology and molecular methods. Infected horses remain viraemic carriers for life and, with very
rare exceptions, yield a positive serological test result. Antibody response usually persists and
antibody-positive animals, older than 6–8 months, are identified as virus carriers (below 6–8 months
of age, serological reactions can be due to maternal antibodies; status can be confirmed by
molecular techniques). Infected equids are potential virus reservoirs. Biting flies are mechanical
vectors for the virus in nature.
Identification of the agent: Virus from a horse can be isolated by inoculating suspect blood into a
susceptible horse or on to leukocyte cultures prepared from susceptible horses. Recognition of
infection in horses that have been inoculated experimentally may be made on the basis of clinical
signs, haematological changes and a positive antibody response determined by an
immunodiffusion test or enzyme-linked immunosorbent assay (ELISA) or by molecular techniques.
Successful virus isolation in horse leukocyte cultures is confirmed by the detection of specific EIA
antigen, by immunofluorescence assay, polymerase chain reaction, reverse-transcriptase assay, or
by the inoculation of culture fluids into susceptible horses. Virus isolation is rarely attempted
because of the time, difficulty and expense involved.
Serological tests: Agar gel immunodiffusion (AGID) tests and ELISAs are simple, reliable tests for
the demonstration of EIAV infection. When ELISAs are positive they should be confirmed using the
AGID test. EIA antigens can be prepared from infected tissue cultures or by using recombinant
DNA technology.
Requirements for vaccines and diagnostic biologicals: There are no biological products
currently available.
A. INTRODUCTION
Equine infectious anaemia (EIA) occurs world-wide. The infection, formerly known as swamp fever, is limited to
equids. The disease is characterised by recurrent febrile episodes, thromboctopenia, anaemia, rapid loss of
weight and oedema of the lower parts of the body. If death does not result from one of the acute clinical attacks, a
chronic stage develops and the infection tends to become inapparent. The incubation period is normally 1–
3 weeks, but may be as long as 3 months. In acute cases, lymph nodes, spleen and liver are hyperaemic and
enlarged. Histologically these organs are infiltrated with nests of immature lymphocytes and plasma cells. Kupffer
cells in the liver often contain haemosiderin or erythrocytes. The enlarged spleen may be felt on rectal
examination. Differential diagnoses include equine viral arteritis (Chapter 2.5.10), and other causes of oedema,
fever, or anemia.
Once a horse is infected with EIA virus (EIAV), its blood remains infectious for the remainder of its life. This
means that the horse is a viraemic carrier and can potentially transmit the infection to other horses (Cheevers &
McGuire, 1985). Transmission occurs by transfer of blood from an infected horse. In nature, spread of the virus is
most likely via interrupted feeding of bloodsucking horseflies on a clinically ill horse and then on susceptible
horses. Transmission can also occur by the iatrogenic transfer of blood through the use of contaminated needles.
In utero infection of the fetus may occur (Kemen & Coggins, 1972). The virus titre is higher in horses with clinical
signs and the risk of transmission is higher from these animals than the carrier animals with a lower virus titre.
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B. DIAGNOSTIC TECHNIQUES
Agar gel immunodiffusion (AGID) tests (Coggins et al., 1972) and enzyme-linked immunosorbent assays
(ELISAs) (Suzuki et al., 1982) are accurate, reliable tests for the detection of EIA in horses, except for animals in
the early stages of infection and foals of infected dams. In other rare circumstances, misleading results may occur
when the level of virus circulating in the blood during an acute episode of the disease is sufficient to bind available
antibody, and if initial antibody levels never rise high enough to be detectable (Toma, 1980). Although the ELISA
will detect antibodies somewhat earlier and at lower concentrations than the AGID test, positive ELISAs are
confirmed using the AGID test. This is because false-positive results have been noted with ELISAs. The AGID
test also has the advantage of distinguishing between EIA and non-EIA antigen–antibody reactions by lines of
identity.
The EIAV is a lentivirus in the family Retroviridae, subfamily Orthoretrovirinae. Other members of the lentivirus
genus include: bovine immunodeficiency virus; caprine arthritis encephalitis virus; feline immunodeficiency virus;
human immunodeficiency virus 1; human immunodeficiency virus 2; and maedi/visna virus. Nucleic acid
sequence comparisons have demonstrated a marked relatedness among these viruses.
1.
Identification of the agent
a)
Virus isolation and identification
Virus isolation is usually not necessary to make a diagnosis.
Isolation of the virus from suspect horses may be made by inoculating their blood on to leukocyte cultures
prepared from horses free of infection. Virus production in cultures can be confirmed by detection of specific
EIA antigen by ELISA (Shane et al., 1984), by imunofluorescence assay (Weiland et al., 1982), by molecular
tests or by subinoculation into susceptible horses. Virus isolation is rarely attempted because of the difficulty
of growing horse leukocyte cultures.
When the exact status of infection of a horse cannot be ascertained, the inoculation of a susceptible horse
with suspect blood should be employed. In this case a horse that has previously been tested for antibody
and shown to be negative is given an immediate blood transfusion from the suspect horse, and its antibody
status and clinical condition are monitored for at least 45 days. Usually, 1–25 ml of whole blood given
intravenously is sufficient to demonstrate infection, but in rare cases it may be necessary to use a larger
volume of blood (250 ml) or washed leukocytes from such a volume (Coggins & Kemen, 1976).
b)
Polymerase chain reaction
A nested polymerase chain reaction (PCR) assay to detect EIA proviral DNA from the peripheral blood of
horses has been described (Nagarajan & Simard, 2001). The nested PCR method is based on primer
sequences from the gag region of the proviral genome. It has proven to be a sensitive technique to detect
field strains of EAV in white blood cells of EIA infected horses; the lower limit of detection is typically around
10 genomic copies of the target DNA (Nagarajan & Simard, 2001; 2007). A real-time reverse-transcriptase
PCR assay has also been described (Cook et al., 2002). To confirm the results of these very sensitive
assays, it is recommended that duplicate samples of each diagnostic specimen be processed. Because of
the risk of cross contamination, it is also important that proper procedures are followed. Methods to insure
the validity of PCR testing are discussed in detail in Chapter 1.1.5 Principles and methods of validation of
diagnostic assays for infectious diseases.
The following are some of the circumstances where the PCR assay maybe used for the detection of EIAV
infection in horses:
•
Conflicting results on serologic tests;
•
Suspected infection but negative or questionable serologic results;
•
Complementary test to serology for the confirmation of positive results;
•
Confirmation of early infection, before serum antibodies to EIAV develop;
•
Ensuring that horses that are to be used for antiserum or vaccine production or as blood donors are
free of EIAV;
•
Confirmation of the status of a foal from an infected mare.
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2.
Serological tests
Due to the persistence of EIAV in infected equids, detection of serum antibody to EIAV confirms the diagnosis of
EIAV infection.
a)
Agar gel immunodiffusion test (the prescribed test for international trade)
Precipitating antibody is rapidly produced as a result of EIA infection, and can be detected by the AGID test.
Specific reactions are indicated by precipitin lines between the EIA antigen and the test serum and
confirmed by their identity with the reaction between the antigen and the positive standard serum. Horses in
the first 2–3 weeks after infection will usually give negative serological reactions. In rare cases the postinfection time prior to the appearance of detectable antibody may extend up to 60 days.
Reagents for AGID are available commercially from several companies. Alternately, AGID antigen and
reference serum may be prepared as described below.
•
Preparation of antigen
Specific EIA antigen may be prepared from the spleen of acutely infected horses (Coggins et al., 1973), from
infected equine tissue culture (Malmquist et al., 1973), from a persistently infected canine thymus cell line
(Bouillant et al., 1986), or from proteins expressed in bacteria or baculovirus using the recombinant DNA
technique (Archambault et al., 1989; Kong et al., 1997). Preparation from infected cultures or from
recombinant DNA techniques gives a more uniform result than the use of spleen cells and allows for better
standardisation of reagents.
To obtain a satisfactory antigen from spleen, a horse must be infected with a highly virulent strain of EIAV.
The resulting incubation period should be 5–7 days, and the spleen should be collected 9 days after
inoculation, when the virus titre is at its peak and before any detectable amount of precipitating antibody is
produced. Undiluted spleen pulp is used in the immunodiffusion test as antigen (Coggins et al., 1973).
Extraction of antigen from the spleen with a saline solution and concentration with ammonium sulphate does
not give as satisfactory an antigen as selection of a spleen with a very high titre of EIA antigen.
Alternatively, equine fetal kidney or dermal cells or canine thymus cells are infected with a strain of EIAV
adapted to grow in tissue culture (American Type Culture Collection). Virus is collected from cultures by
precipitation with 8% polyethylene glycol or by pelleting by ultracentrifugation. The diagnostic antigen, p26,
is released from the virus by treatment with detergent or ether (Malmquist et al., 1973). EIAV core proteins,
expressed in bacteria or baculovirus, are commercially available and find practical use as high quality
antigens for serological diagnosis.
The p26 is an internal structural protein of the virus that is coded for by the gag gene. The p26 is more
antigenically stable among EIAV strains than the virion glycoproteins gp45 and gp90 (Montelaro et al.,
1984). There is evidence of strain variation in the p26 amino acid sequence; however there is no evidence to
indicate that this variation influences any of the serological diagnostic tests (Zhang et al., 1999).
•
Preparation of standard antiserum
A known positive antiserum may be collected from a horse previously infected with EIAV. This serum should
yield a single dense precipitation line that is specific for EIA, as demonstrated by a reaction of identity with a
known standard serum. It is essential to balance the antigen and antibody concentrations in order to ensure
the optimal sensitivity of the test. Reagent concentrations should be adjusted to form a narrow precipitation
line approximately equidistant between the two wells containing antigen and serum.
862
•
Test procedure (Association Française de Normalisation [AFNOR], 2000; Coggins et al., 1973; PEARSON
& COGGINS, 1979)
i)
Immunodiffusion reactions are carried out in a layer of agar in Petri dishes. For Petri dishes that are
100 mm in diameter, 15–17 ml of 1% Noble agar in 0.145 M borate buffer (9 g H3BO3, plus 2 g NaOH
per litre), pH 8.6 (± 0.2) is used. Six wells are punched out of the agar surrounding a centre well of the
same diameter. The wells are 5.3 mm in diameter and 2.4 mm apart. Each well must contain the same
volume of reagent.
ii)
The antigen is placed in the central well and the standard antiserum is placed in alternate exterior
wells. Serum samples for testing are placed in the remaining three wells.
iii)
The dishes are maintained at room temperature in a humid environment.
iv)
After 24–48 hours the precipitation reactions are examined over a narrow beam of intense, oblique light
and against a black background. The reference lines should be clearly visible at 24 hours, and at that
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Chapter 2.5.6. — Equine infectious anaemia
time any test sera that are strongly positive may also have formed lines of identity with those between
the standard reagents. A weakly positive reaction may take 48 hours to form and is indicated by a
slight bending of the standard serum precipitation line between the antigen well and the test serum
well. Sera with high precipitating antibody titres may form broader precipitin bands that tend to be
diffuse. Such reactions can be confirmed as specific for EIA by dilution at 1/2 or 1/4 prior to retesting;
these then give a more distinct line of identity. Sera devoid of EIA antibody will not form precipitation
lines and will have no effect on the reaction lines of the standard reagents.
v)
b)
Interpretation of the results: Horses that are in the early stages of an infection may not give a positive
serological reaction in an AGID test. Such animals should be bled again after 3–4 weeks. In order to
make a diagnosis in a young foal, it may be necessary to determine the antibody status of the dam. If
the mare passes EIA antibody to the foal through colostrum, then a period of 6 months or longer after
birth must be allowed for the maternal antibody to wane. Sequential testing of the foal at monthly
intervals may be useful to observe the decline in maternal antibody. To conclude that the foal is not
infected, a negative result must be obtained (following an initial positive result) at least 2 months after
separating the foal from contact with the EIA positive mare or any other positive horse. Alternatively
PCR could be performed on the blood of the foal to determine the presence/absence of EIA proivirus.
Enzyme-linked immunosorbent assay
There are four ELISAs that are approved by the United States Department of Agriculture for the diagnosis of
equine infectious anaemia and are available internationally; a competitive ELISA and three non-competitive
ELISAs. The competitive ELISA and two non-competitive ELISAs detect antibody produced against the p26
core protein antigen. The third non-competitive ELISA incorporates both p26 core protein and gp45 (viral
transmembrane protein) antigens. Typical ELISA protocols are used in all tests. If commercial ELISA
materials are not available, a non-competitive ELISA using p26 antigen purified from cell culture material
may be employed (Shane et al., 1984).
A positive test result by ELISA should be retested using the AGID test to confirm the diagnosis because
some false-positive results have been noted with the ELISA. The results can also be confirmed by the
immunoblot technique. A standard antiserum for immunodiffusion, which contains the minimum amount of
antibody that should be detected by laboratories, is available from the OIE Reference Laboratories (see
Table given in Part 4 of this Terrestrial Manual). Uniform methods for EIA control have been published
(United States Department of Agriculture [USDA], 2002).
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
No biological products are available currently.
REFERENCES
ASSOCIATION FRANÇAISE DE NORMALISATION (AFNOR) (2000). Animal Health Analysis Methods. Detection of
Antibodies against Equine Infectious Anaemia by the Agar Gel Immunodiffusion Test. NF U 47-002. AFNOR, 11
avenue Francis de Pressensé, 93571 Saint-Denis La Plaine Cedex, France.
ARCHAMBAULT D., WANG Z., LACAL J.C., GAZIT A., YANIV A., DAHLBERG J.E. & TRONICK S.R. (1989). Development of
an enzyme-linked immunosorbent assay for equine infectious anaemia virus detection using recombinant
Pr55gag. J. Clin. Microbiol., 27, 1167–1173.
BOUILLANT A.M.P., NELSON K., RUCKERBAUER C.M., SAMAGH B.S. & HARE W.C.D. (1986). The persistent infection of
a canine thymus cell line by equine infectious anaemia virus and preliminary data on the production of viral
antigens. J. Virol. Methods, 13, 309–321.
CHEEVERS W.M. & MCGUIRE T.C. (1985). Equine infectious anaemia virus; immunopathogenesis and persistence.
Rev. Infect. Dis., 7, 83–88.
COGGINS L. & KEMEN M.J. (1976). Inapparent carriers of equine infectious anaemia (EIA) virus. In: Proceedings of
the IVth International Conference on Equine Infectious Diseases. University of Kentucky, Lexington, Kentucky,
USA, 14–22.
COGGINS L., NORCROSS N.L. & KEMEN M.J. (1973). The technique and application of the immunodiffusion test for
equine infectious anaemia. Equine Infect. Dis., III, 177–186.
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Chapter 2.5.6. — Equine infectious anaemia
COGGINS L., NORCROSS N.L. & NUSBAUM S.R. (1972). Diagnosis of equine infectious anaemia by immunodiffusion
test. Am. J. Vet. Res., 33, 11–18.
COOK R.F., COOK S.J., LI F.L., MONTELARO R.C., & ISSEL C.J. (2002). Development of a multiplex real-time reverse
transcriptase-polymerase chain reaction for equine infectious anemia virus (EIAV). Virol Methods, 105, 171–179.
KEMEN M.J. & COGGINS L. (1972). Equine infectious anaemia: transmission from infected mares to foals. J. Am.
Vet. Med. Assoc., 161, 496–499.
KONG X. K., PANG H., SUGIURA T., SENTSUI H., ONODERA T., MATSUMOTO Y. & AKASHI H. (1997). Application of equine
infectious anaemia virus core proteins produced in a Baculovirus expression system, to serological diagnosis.
Microbiol. Immunol., 41, 975–980.
MALMQUIST W.A., BARNETT D. & BECVAR C.S. (1973). Production of equine infectious anaemia antigen in a
persistently infected cell line. Arch. Gesamte Virusforsch., 42, 361–370.
MONTELARO R.C., PAREKH B., ORREGO A. & ISSEL C.J. (1984). Antigenic variation during persistent infection by
equine infectious anaemia, a retrovirus. J. Biol. Chem., 16, 10539–10544.
NAGARAJAN M.M. & SIMARD C. (2001). Detection of horses infected naturally with equine infectious anemia virus by
nested polymerase chain reaction. J. Virol. Methods, 94, 97–109.
NAGARAJAN M.M. & SIMARD C. (2007). Gag genomic heterogeneity of equine infectious anemia virus (EIAV) in
naturally infected horses in Canada: implication on EIA diagnosis and peptide-based vaccine development. Virus
Res., 129, 228–235.
PEARSON J.E. & COGGINS L. (1979). Protocol for the immunodiffusion (Coggins) test for equine infectious anaemia.
Proc. Am. Assoc. Vet. Lab. Diagnosticians, 22, 449–462.
SHANE B.S., ISSEL C.J. & MONTELARO R.C. (1984). Enzyme-linked immunosorbent assay for detection of equine
infectious anemia virus p26 antigen and antibody. J. Clin. Microbiol., 19, 351–355.
SUZUKI T., UEDA S. & SAMEJINA T. (1982). Enzyme-linked immunosorbent assay for diagnosis of equine infectious
anaemia. Vet. Microbiol., 7, 307–316.
TOMA B. (1980). Réponse sérologique négative persistante chez une jument infectée. Rec. Med. Vet., 156, 55–63.
UNITED STATES DEPARTMENT OF AGRICULTURE (USDA) (2002). Equine Infectious Anemia Uniform Methods and
Rules. Animal and Plant Health Inspection Service, USDA:
http://www.aphis.usda.gov/animal_health/animal_diseases/eia/
WEILAND F., MATHEKA H.D. & BOHM H.O. (1982). Equine infectious anaemia: detection of antibodies using an
immunofluorescence test. Res. Vet. Sci., 33, 347–350.
ZHANG W., AUYONG D.B., OAKS J.L. & MCGUIRE T.C. (1999). Natural variation of equine infectious anemia virus
Gag-protein cytotoxic T lymphocyte epitopes. Virology, 261, 242–252.
*
* *
NB: There are OIE Reference Laboratories for Equine infectious anaemia:
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests and reagents for equine infectious anaemia
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CHAPTER 2.5.7.
EQUINE INFLUENZA
SUMMARY
Equine influenza is an acute respiratory infection of horses, donkeys and mules caused by two distinct
subtypes (H7N7, formerly equi-1, and H3N8, formerly equi-2) of influenza A virus within the genus
Influenzavirus A of the family Orthomyxoviridae. Viruses of the H7N7 subtype have not been isolated
since the late 1970s. Equine influenza viruses of both subtypes are considered to be of avian ancestry
and highly pathogenic avian H5N1 has recently been associated with an outbreak of respiratory
disease in donkeys in Egypt. In fully susceptible equidae, clinical signs include pyrexia and a harsh
dry cough followed by a mucopurulent nasal discharge. In partially immune vaccinated animals, one
or more of these signs may be absent. Vaccinated infected horses can still shed the virus and serve
as a source of virus to their cohorts. Characteristically, influenza spreads rapidly in a susceptible
population. The disease is endemic in many countries with substantial equine populations. In recent
years, infection has been introduced into Australia and re-introduced into South Africa and Japan; to
date New Zealand and Iceland are reported to be free of equine influenza virus.
While normally confined to equidae, equine H3N8 influenza has crossed the species barrier to
dogs. Extensive infection of dogs has been reported in North America where it normally produces
mild fever and coughing but can cause fatal pneumonia. While equine influenza has not been
shown to cause disease in humans, serological evidence of infection has been described primarily
in individuals with an occupational exposure to the virus. During 2004–2006 influenza surveillance
in central China (People’s Rep. of) two equine H3N8 influenza viruses were also isolated from pigs.
Identification of the agent: Embryonated hens’ eggs and/or cell cultures can be used for virus
isolation from nasopharyngeal swabs or nasal and tracheal washes. Isolates should always be sent
immediately to an OIE Reference Laboratory. Infection may also be demonstrated by detection of
viral nucleic acid or antigen in respiratory secretions using the reverse-transcription polymerase
chain reaction (RT-PCR) or an antigen-capture enzyme-linked immunosorbent assay (ELISA),
respectively.
Serological tests: Diagnosis of influenza virus infections is usually only accomplished by tests on
paired sera; the first sample should be taken as soon as possible after the onset of clinical signs
and the second approximately 2 weeks later. Antibody levels are determined by haemagglutination
inhibition (HI) or single radial haemolysis (SRH).
Requirements for vaccines and diagnostic biologicals: Spread of infection and severity of
disease may be reduced by the use of potent inactivated equine influenza vaccines containing
epidemiologically relevant virus strains. Inactivated equine influenza vaccines contain whole viruses
or their subunits. The vaccine viruses are propagated in embryonated hens’ eggs or tissue culture,
concentrated, and purified before inactivation with agents such as formalin or beta-propiolactone.
Inactivated vaccines provide protection by inducing humoral antibody to the haemagglutinin protein.
Responses are generally short-lived and multiple doses are required to maintain protective levels of
antibody. An adjuvant is usually required to stimulate durable protective levels of antibody. Live
attenuated virus and viral vectored vaccines have been licensed in some countries.
Vaccine breakdown has been attributed to inadequate vaccine potency, inappropriate vaccination
schedules, and outdated vaccine viruses that are compromised as a result of antigenic drift. An invitro potency test (single radial diffusion) can be used for in-process testing of the antigenic content
of inactivated products before addition of an adjuvant. In process testing of live and vectored
vaccines relies on titration of infectious virus. A surveillance programme is underway to monitor
antigenic drift among equine influenza viruses and to provide information on strain selection for
vaccines.
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Chapter 2.5.7. — Equine influenza
A. INTRODUCTION
Equine influenza is caused by two subtypes: H7N7 (formerly subtype 1) and H3N8 (formerly subtype 2) of
influenza A viruses (genus Influenzavirus A of the family Orthomyxoviridae); however there have been very few
reports of H7N7 subtype virus infections in the last 30 years (Webster, 1993).
In fully susceptible equidae, clinical signs include pyrexia, nasal discharge and a harsh dry cough; pneumonia in
young foals and donkeys and encephalitis in horses have been described as rare events (Daly et al., 2006;
Gerber, 1970). Clinical signs associated with infection in dogs also include fever and a cough; occasionally
infection results in suppurative bronchopneumonia and peracute death (Crawford et al., 2005). Characteristically,
influenza spreads rapidly in a susceptible population. The virus is spread by the respiratory route, and indirectly
by contaminated personnel, vehicles and fomites. The incubation period in susceptible horses may be less than
24 hours. In partially immune vaccinated animals the incubation period may be extended, one or more clinical
signs may be absent and spread of the disease may be limited. This makes clinical diagnosis of equine influenza
more difficult as other viral diseases, such as equine herpesvirus-associated respiratory disease, may clinically
resemble a mild form of influenza. Horses infected with equine influenza virus become susceptible to secondary
bacterial infection and may develop mucopurulent nasal discharge, which can lead to diagnosis of bacterial
disease with the underlying cause being overlooked.
Equine influenza viruses are believed to be of avian ancestry, and more recent transmission of avian viruses to
horses and donkeys has been recorded. The sequence analysis of an H3N8 virus isolated in 1989 from horses
during a limited influenza epidemic in North Eastern China (People’s Rep. of) established that the virus was more
closely related to avian influenza viruses than to equine influenza viruses (Guo et al., 1992). Recently, avian
H5N1 was associated with respiratory disease of donkeys in Egypt (Abdel-Moneim et al., 2010).
Equine influenza viruses have the potential to cross species barriers and have been associated with respiratory
disease in dogs primarily in North America (Crawford et al., 2005). Isolated outbreaks of equine influenza have
also occurred in dogs within the UK but the virus has not become established in the canine population. Close
contact with infected horses was thought to be involved in each outbreak in the UK. Equine influenza viruses have
also been isolated from pigs in central China (People’s Rep. of) (Tu et al., 2009). Despite the occasional
identification of seropositive persons with occupational exposure there is currently little evidence of zoonotic
infection of people with equine influenza (Alexander & Brown, 2000).
In endemic countries the economic losses due to equine influenza can be minimised by vaccination and many
racing authorities and equestrian bodies have mandatory vaccination policies. Vaccination does not produce
sterile immunity; vaccinated horses may shed virus and contribute silently to the spread of EI. Appropriate risk
management strategies to deal with this possibility should be developed.
B. DIAGNOSTIC TECHNIQUES
Laboratory diagnosis of acute equine influenza virus infections is based on virus detection in nasal swabs
collected from horses with acute respiratory illness. Alternatively, the demonstration of a serological response to
infection may be attempted with paired serum samples. Ideally, both methods are used. Equine influenza virus
may be isolated in embryonated hens’ eggs or cell culture. Infection may also be demonstrated by detection of
viral antigen in respiratory secretions using an antigen capture enzyme-linked immunosorbent assay (ELISA) or of
viral genome using reverse-transcription polymerase chain reaction (RT-PCR) assays. All influenza viruses are
highly contagious for susceptible hosts and care must therefore be taken during the handling of infected eggs or
cultures to avoid accidental cross-contamination. Standard strains should not be propagated in the diagnostic
laboratory, at least never at the same time or in the same place where diagnostic samples are being processed.
All working areas must be efficiently disinfected before and after virus manipulations, which should preferably be
conducted within biohazard containment level 2 and class II safety cabinets.
It is important to obtain samples as soon as possible after the onset of clinical signs, preferably within 3–5 days.
These samples include nasopharyngeal swabs and nasal or tracheal washings, the latter taken by endoscopy.
Swabs may consist of absorbent cotton wool sponge/gauze on wire, and should be long enough to be passed via
the ventral meatus into the nasopharynx. Swabs should be transferred to a tube containing transport medium
immediately after use. This medium consists of phosphate buffered saline (PBS) containing either 40% glycerol or
2% tryptose phosphate broth with 2% antibiotic solution (penicillin [10,000 units], streptomycin [10,000 units] in
sterile distilled water [100 ml]), and 2% fungizone (250 mg/ml stock). If the samples are to be inoculated within 1–
2 days they may be held at 4°C, but, if kept for longer, they should be stored at –70°C or below. Samples should
be kept cool during transport to the laboratory.
Sample processing should follow the quality procedures outlined in Chapter 1.1.4 Quality management in veterinary
testing laboratories, taking measures to prevent cross contamination. The liquid is expelled from the swab by squeezing
with forceps and the swab is then disposed of suitably. Further antibiotics may be added if samples appear to be heavily
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contaminated with bacteria. The remainder of the fluid is stored at –70°C. Samples treated with antibiotics are allowed
to stand on ice for 30–60 minutes and are then centrifuged at 1500 g for 15 minutes to remove bacteria and debris; the
supernatant fluids are used for inoculation. The remainder of the fluid is stored at –70°C. Filtration of samples is not
advised as influenza virus may adsorb on to the filter and be lost from the sample.
1.
Identification of the agent
Isolation of infectious virus may be carried out in embryonated hens’ eggs or cell cultures. Traditionally, eggs
have been preferred for isolation of equine influenza as many clinical isolates do not grow well in cells without
serial passage. Comparison of H3N8 viruses isolated in eggs and Madin–Darby canine kidney (MDCK) cells
indicated that MDCK cells are capable of selecting variant viruses that are not representative of the predominant
virus in clinical specimens (Ilobi et al., 1994). However, some viruses have been successfully isolated in MDCK
cells but not in eggs and selection of variants has also occurred as a result of culture in eggs (Oxburgh &
Klingborn, 1999). Ideally, isolation should be attempted using both substrates.
RT-PCR and real-time RT-PCR assays are being widely used in diagnostic laboratories as a more sensitive
alternative to virus isolation (Quinlivan et al., 2005). Influenza virus antigen in nasal secretions may also be
detected directly by a sensitive antigen-capture ELISA for the H3N8 virus using a monoclonal antibody (MAb)
against the equine influenza virus nucleoprotein (Livesay et al., 1993).This assay is not commercially available,
other than as a diagnostic service, but commercial self-contained kits for detecting human influenza are available
and have been shown to detect equine influenza antigen (Chambers et al., 1994; Yamanaka et al., 2008). This
approach is less sensitive than RT-PCR but provides a rapid result on which management decisions may be
based. It should not be used to the exclusion of virus isolation. It is essential that new viruses be isolated and sent
to reference laboratories for characterisation as part of the surveillance programme to monitor antigenic drift and
emergence of new viruses and to provide isolates for inclusion in updated vaccines. Positive RT-PCR and ELISA
results are useful in the selection of samples for virus isolation attempts if resources are limited, or for the
selection of specimens to be sent to a reference laboratory for virus isolation and characterisation.
a)
Virus isolation in embryonated hens’ eggs
Fertile eggs are set in a humid incubator at 37–38°C and turned twice daily; after 10–11 days, they are
examined by candling and live embryonated eggs are selected for use. The area above the air sac is
cleansed with alcohol and a small hole is made through the shell. Inoculum may be introduced into the
amniotic or the allantoic cavity. Several eggs/sample are inoculated (0.1 ml) in the amniotic cavity with no
additional dilution of the sample (sample may also be diluted 1/10 and 1/100 in PBS containing antibiotics).
The syringe is withdrawn approximately 1 cm and a further 0.1 ml is inoculated into the allantoic cavity.
Alternatively, many laboratories opt to inoculate into the allantoic cavity alone through a second hole drilled
just below the line of the air sac. The hole(s) is/are sealed with wax or Sellotape, and the eggs are incubated
at 34–35°C for 3 days. The embryos that die within 24 hours following inoculation should be discarded. The
eggs that contain embryos that die more than 24 hours after inoculation or contain live embryos after 3 days
are examined for the presence of EI virus.
The eggs are transferred to 4°C for 4 hours or overnight to kill the embryos and to reduce bleeding at
harvest. The shells are disinfected, and the amniotic and/or allantoic fluid is harvested by pipette, each
harvest being kept separate. These are tested for haemagglutination (HA) activity by mixing twofold dilutions
of the harvested fluid in equal volumes (0.025 ml) with chicken red blood cells (RBCs) (0.5% [v/v] packed
cells in PBS) in V- or U-bottomed microtitre plates or 0.4% guinea-pig RBCs (0.4% [v/v] packed cells in PBS)
in V- or U-bottomed plates. Plates are incubated for approximately 30 minutes preferably at 4°C to prevent
neuraminidase activity. If chicken RBCs are used, the plates may be read by tilting to 70° so that nonagglutinated cells ‘stream’ to the bottom of the well. Non-agglutinated guinea-pig cells appear as a button at
the bottom of the well and may take longer to settle. If there is no HA activity, aliquots of each harvest are
pooled and passaged into further eggs. All HA positive samples are divided into aliquots and stored at
–70°C; one aliquot is titrated for HA immediately. The HA titre is the reciprocal of the greatest dilution to
show agglutination. If the HA titre is 1/16 or more, the isolate is characterised immediately. If titres are low,
positive samples should be passaged. Care should be taken to avoid generation of defective interfering
particles by prediluting the inoculum 1/10, 1/100, 1/1,000. Positive samples arising from the highest dilution
should be selected as stocks for storage. It may be necessary to undertake as many as five passages to
isolate the virus, particularly from vaccinated horses. If virus has not been recovered by the fifth passage,
further passages are unlikely to be successful.
b)
Virus isolation in cell cultures
Cultures of the MDCK cell line (MDCK, ATCC CCL34) may be used to isolate equine influenza viruses. The
cells are grown to confluence in tubes and then infected in triplicate with 0.25–0.5 ml of each sample,
processed as described above. Prior to inoculation, the cell monolayer is washed at least once with tissue
culture medium containing trypsin (2 µg/ml) without serum. The cultures are maintained with serum-free
medium containing 0.5–2 µg/ml trypsin (treated with TPCK [L-1-tosylamine-2-phenylethyl chloromethyl
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Chapter 2.5.7. — Equine influenza
ketone] to remove chymotrypsin, available pretreated, e.g. from Sigma), and examined daily for evidence of
cytopathic effects (CPE). If positive, or after 7 days in any case, the supernatant fluids are tested for HA.
Fluids with titres of ≥1/16 are characterised immediately. Negative fluids and those with titres <1/16 are
repassaged up to five passages.
Alternatively, the cells are screened for evidence of haemadsorption (HAD). This procedure detects
expression of viral antigens at the cell surface. The medium is removed from the cultures and the tubes are
washed with PBS. One or two drops of a 50% suspension of chicken or guinea-pig RBCs are added, the
tubes are rotated carefully, and kept at room temperature (23°C ± 2°C) for 30 minutes. Unbound RBCs are
washed off with PBS, and the cultures are examined microscopically for evidence of HAD.
c)
Haemagglutinin subtyping
The HA subtype of new isolates of equine influenza viruses is best determined by haemagglutination
inhibition (HI; Section B.2.a) using H7N7- and H3N8-specific antisera. Isolates may first be treated with
Tween 80/ether, which destroys viral infectivity and reduces the risk of cross-contamination. In the case of
H3N8 viruses particularly, this treatment enhances the HA activity (John & Fulginiti, 1966). However,
treatment with Tween 80/ether also decreases specificity and may increase the variability of the results
obtained. Standard antigens must be titrated in parallel with tests to identify viruses and should include
H7N7 strains (e.g. A/eq/Prague/56, A/eq/Newmarket/77) and H3N8 strains (e.g. A/eq/Newmarket/2/93, and
A/eq/South Africa/4/03 and A/eq/Richmond/1/07). Virus strains may be obtained from OIE Reference
Laboratories (see Table given in Part 4 of this Terrestrial Manual). Additionally, recent isolates from the
same geographical area should be included if available. The standard antigens should be treated with
Tween 80/ether to avoid cross-contamination. Test antigens and standard antigens are always back-titrated
to confirm their antigen content.
New isolates of equine influenza viruses may be further characterised by HI using strain-specific antisera.
The species in which antibodies are raised will influence the cross-reactivity of the antiserum, with ferrets
providing the most strain-specific antibody (Mumford, 1992). The specificity and cross reactivity of the sera
are also influenced by the immunisation schedule. Sera obtained 3 weeks after a single antigen application
are considered to be most discriminative.
All isolates should be sent immediately to an International Reference Laboratory designated by OIE or the
World Health Organization (WHO) for inclusion in the strain surveillance programme to monitor antigenic
drift and emergence of new viruses.
d)
Neuraminidase subtyping
Subtyping of neuraminidase requires specific antisera and no routine technique is available. Subtyping can
also be done using specific PCR primers.
e)
Polymerase chain reaction
RT-PCR assays both conventional and real-time, are increasingly used for the detection of equine influenza
genome in nasal secretions. Real-time Light Cycler RT-PCR using primers M52C (5’-CTT-CTA-ACC-GAGGTC-GAA-ACG-3’) and M253R (5’-AGG-GCA-TTT-TGG-ACA-AAG-CGT-CTA-3’) designed to amplify a
244 base pair amplicon from nucleotides 32 to 276 of the matrix gene, which is highly conserved across
influenza A viruses, has been shown to be more sensitive for the detection of positive samples than virus
culture in eggs or detection of nucleoprotein using a kit for the detection of human influenza (Tu et al., 2009).
For several years this assay has been used effectively for diagnosis and surveillance by one of the OIE
reference laboratories. However it has not been validated by interlaboratory comparisons at international
level. A probe-based real-time RT-PCR assay based on the matrix gene and developed for the detection of
a wide range of influenza type A strains including avian H5N1, was combined with an automated DNA
extraction system to establish a high throughput assay used in the mass screening of horses during the
eradication programme in Australia in 2007 (Foord et al., 2009).This type of technology is constantly
evolving. TaqMan® real-time RT-PCR assays specific for equine-2 influenza (H3N8) and equine-1 (H7N7)
viruses have recently been described and a commercial RT-PCR kit for the detection of equine influenza has
become available (Lu et al., 2009). These RT-PCR assays have not yet been validated in line with the OIE
Validation Template.
Although the genetic sequence of isolates can also be derived from PCR assays it remains essential to
isolate infectious virus in order to examine the antigenic properties of new isolates and evaluate antigenic
drift in the field.
2.
Serological tests
Influenza infections can be detected by performing serological tests on paired sera to show a rise in specific
antibody. These tests should be carried out whether virus isolation has been attempted or not. They are robust
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Chapter 2.5.7. — Equine influenza
and may prove positive in the absence of virus isolation. Two simple methods exist, HI and single radial
haemolysis (SRH), each equally efficient and widely used. The complement fixation (CF) test can also be applied,
but is not in general use. The paired serum samples, i.e. the acute and convalescent samples, should be tested
together to minimise the impact of inter-assay variability. The standard antigens are described above (Section
B.1.c). If available, isolates from recent cases should be included. Freeze-dried post-infection equine antisera to
A/eq/Newmarket/77 (H7N7), A/eq/Newmarket/1/93 (American lineage H3N8), A/eq/Newmarket/2/93 (European
lineage H3N8), A/eq/South Africa/4/03 (Florida Clade 1, sublineage of American lineage) and an influenzanegative equine serum, are available from the European Directorate for the Quality of Medicines (EDQM)1. These
sera have been assigned SRH values through an international collaborative study and can be used as primary
reference sera for the assay (Daly et al., 2007; Mumford, 2000).
a)
Haemagglutination inhibition test
The antigen is first treated with Tween 80/ether in order to increase the sensitivity of the test, particularly for
H3N8 viruses. The test is best done in microtitre plates using the appropriate dilution equipment. A
macrotest may be used, for which antigen is diluted to a final HA titre of 1/8 per well and the volumes for
PBS, sera and antigen are 0.5 ml. Sera are pretreated to remove nonspecific haemagglutinins, then
inactivated at 56°C for 30 minutes. Pretreatments include the use of one of the following: (a) kaolin and
RBCs absorption, (b) potassium periodate, or (c) Vibrio cholerae receptor-destroying enzyme. Potassium
periodate or V. cholerae receptor-destroying enzyme is the treatment of choice. The treated sera are diluted
in PBS, a standard dose of antigen is added (HA titre of 1/4 per well for microtitration assay), and these are
kept at room temperature (23°C ± 2°C) for 30 minutes. After gentle mixing, RBCs are added and the test is
read 30 minutes later. The HI titres are read as the highest dilution of serum giving complete inhibition of
agglutination. Either chicken RBCs (1% [v/v] packed cells) in V-bottomed microtitre plates or guinea-pig
RBCs (0.5% [v/v] packed cells) in V- or U-bottomed plates may be used. If chicken RBCs are used, the
plates may read by tilting to 70° so that non-agglutinated cells ‘stream’ to the bottom of the well. Nonagglutinated guinea-pig cells appear as a ‘button’ in the bottom of the well and may take longer to settle. The
HI titre is the reciprocal of the greatest dilution showing complete inhibition of agglutination. Currently, a cutoff point for positive samples has not been determined for the HI test and thus, low titres should be
investigated further. Titre increases of fourfold or more between paired sera indicate recent infection.
1
•
Tween 80/ether treatment of the virus
i)
To 39.5 ml of infective allantoic fluid, add 0.5 ml of a 10% (v/v) suspension of Tween 80 in PBS to give
a 0.125% (v/v) concentration of Tween 80.
ii)
After mixing gently at room temperature for 5 minutes, add 20 ml of diethyl ether to give a final
concentration of 33.3% by volume, and mix the suspension well at 4°C for 15 minutes.
iii)
After allowing the layers to separate by standing, remove the aqueous layer containing the disrupted
virus particles to a glass bottle with a loose lid and allow the excess ether to evaporate off overnight
(John & Fulginiti, 1966). Safety precautions while handling ether must be strictly observed and work
should be confined to a fume hood.
iv)
Store treated virus in aliquots at –70°C.
•
Titration of haemagglutination
i)
Add 25 µl of PBS to all wells in a row of a microtitre plate.
ii)
Add 25 µl of virus to first well then generate a twofold dilution series across the plate leaving the last
well as a negative control.
iii)
Add an extra 25 µl of PBS to all wells.
iv)
Add 50 µl of RBCs to all wells. Leave at room temperature or at 4°C (particularly if ambient
temperatures are high), for 30 minutes. The HA titre is taken as the last virus dilution giving partial HA.
•
Potassium periodate pretreatment of sera
i)
Mix one volume (150 µl) of serum with two volumes (300 µl) of freshly prepared 0.016 M potassium
periodate (0.38 g in 100 ml PBS), and leave at 22°C (± 2°C) for 15 minutes.
ii)
Add a further one volume of 3% glycerol in PBS to neutralise any excess periodate solution, mix and
leave at room temperature (23°C ± 2°C) for 15 minutes.
iii)
Inactivate in a 56°C water bath for 30 minutes.
Headquarters: EDQM - Council of Europe, 7 allée Kastner, CS 30026, F-67081 Strasbourg, France.
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Chapter 2.5.7. — Equine influenza
b)
•
Test procedure
i)
Dispense 25 µl of PBS to all wells of a microtitre plate.
ii)
Add serum (25 µl) to the first well of a row of 12, then generate twofold serial dilutions (1/8 to at least
1/512, allowing for dilution of 1/4 from treatment of serum), leaving the last well as a control.
iii)
Dilute the antigen to give a dose of 4 HA units (4 × minimum agglutinating dose, i.e. titre/4).
iv)
Add 25 µl to each well, and incubate at room temperature (22°C ± 2°C) for 30 minutes.
v)
Add 50 µl of RBCs to each well. Leave at room temperature or at 4°C (particularly if ambient
temperatures are high), for 30 minutes.
vi)
The plates may be read by tilting to 70° so that non-agglutinated cells ‘stream’ to the bottom of the well.
No agglutination is recorded as a positive result.
Single radial haemolysis
In this test, viral antigens are coupled to fixed RBCs that are suspended in agarose containing guinea-pig
complement (C’). Wells are punched in the agarose and filled with test sera. Influenza antibodies and C’ lyse
the antigen-coated RBCs, resulting in a clear, haemolytic zone around the well; the size of this zone is
directly proportional to the level of strain-specific antibody in the serum sample (Morley et al., 1995).
Special immunodiffusion plates (MP Biomedical) may be used for the assay, but simple Petri dishes are also
suitable. Sheep RBCs collected into Alsever’s solution are washed three times. The C’ can be obtained
commercially, or normal guinea-pig serum can be used. The viral antigens are egg-grown stocks or purified
preparations; the strains used are the same as for the HI tests. The viruses are coupled to RBCs by
potassium periodate or by chromic chloride. The coupled antigen/RBCs preparations are mixed with C’,
together with a 1% solution of agarose (low melting grade) in PBS. Care must be taken to ensure that the
temperature is not allowed to rise above 42°C at any time. The mixture is poured into plates and left to set.
Wells of 3 mm in diameter and 12 mm apart are punched in the solidified agarose, at least 6 mm from the
edge of the plates. Such plates may be stored at 4°C for 12 weeks. Plates are prepared for each antigen.
Sera are inactivated at 56°C for 30 minutes, but no further treatment is necessary. Paired sera should be
assayed on the same plate. As a minimum, a subtype-specific antiserum should be included as a control
serum in one well on each plate. All sera are tested in a control plate containing all components except virus
to check for nonspecific lysis. Alternatively, an unrelated virus, such as A/PR/8/34 (H1N1), may be used in
the control plate. Sera that show haemolytic activity for sheep RBCs must be pre-absorbed with sheep
RBCs. Zones of lysis should be clear and not hazy or translucent. All clear zones should be measured and
the area of haemolysis calculated.
•
Preparation of reagents
i)
Saline/HEPES: 0.85% NaCl (4.25 g/500 ml); 0.05 M HEPES (N-2-hydroxyethylpiperazine, N-2ethanesulphonic acid; 5.95 g/500 ml); and 0.02% sodium azide. Make to pH 6.5 with NaOH.
ii)
Saline/HEPES/BSA: as saline/HEPES with 0.2% (w/v) bovine serum albumin (BSA).
iii)
CrCl3 stock solution (2.25 M) 6 g/10 ml: Make fresh 1/400 dilution in 0.85% NaCl for each assay.
iv)
PBS (London)/PBS ‘A’: NaCl (10.00 g); KCl (0.25 g); Na2PO4 (1.45 g); KH2PO4 (0.25 g); and Na azide
(0.20 g). Make up to 1 litre with distilled water.
v)
Agarose in PBS: Place flask containing PBS ‘A’ on a stirrer. Slowly add 10 g agarose to the stirring
solution. Liquefy in a pressure cooker. Dispense into glass bottles for storage at 22°C (± 2°C).
vi)
Virus antigen: Allantoic fluid containing infectious virus is harvested and stored at –70°C. A short
titration curve determines the optimum ratio of virus antigen to RBCs to be used when preparing
sensitised sheep RBCs. The H7N7 influenza strains always produce clear zones; the H3N8 strains
sometimes produce hazy zones, in which case it is necessary to concentrate the virus by
centrifugation.
vii)
Sheep blood: Collect blood into an equal volume of Alsever’s solution and store at 4°C. It may be
necessary to test bleed several sheep, as characteristics of RBCs from individual sheep vary. Keep the
blood for 2 days before use, it may then be usable for up to 3 weeks, providing sterility is maintained.
viii) Complement: Use commercially available guinea-pig complement or collect serum from young guineapigs of 300–350 g body weight and store in small volumes at –70°C. For use, thaw in cold water and
hold at 4°C prior to mixing.
ix)
870
Treatment of sera: Use undiluted sera heat inactivated at 56°C for 30 minutes. Avoid repeated freeze–
thaw cycles.
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Chapter 2.5.7. — Equine influenza
•
Test procedure
i)
Wash sheep RBCs at least three times in saline/HEPES.
ii)
Prepare an appropriate volume of 8% RBCs (v/v packed cells) in saline/HEPES, having first calculated
the number of plates required and allowing 1 ml per 6 × 11 cm immunoplate and 1–2 ml extra.
iii)
Add a predetermined volume2 of virus antigen to the 8% RBCs solution. Hold the mixture at 4°C for
10 minutes. Haemagglutination may be observed.
iv)
SLOWLY add CrCl3 (1/400 in 0.85% NaCl) at half the total volume of virus/RBCs suspension. Hold at
22°C (±2°C) for 5 minutes with occasional mixing.
v)
Sediment the sensitised RBCs by centrifugation at 1500 g for 5 minutes.
vi)
Gently resuspend in saline/HEPES/BSA and centrifuge at 1500 g for 5 minutes.
vii)
Resuspend RBCs to an 8% suspension in PBS ‘A’.
During the sensitisation process, melt the agarose in a pressure cooker. Shortly before use, pipette 7.8 ml
volumes to Universal bottles and retain at 42°C. Check that the agar has cooled to 42°C before use.
•
Preparation of plates
i)
Add 0.9 ml of virus-sensitised sheep RBCs to 7.8 ml of agarose (42°C). Mix quickly, but gently.
ii)
Add 0.3 ml of undiluted guinea-pig serum. Mix again and pour into immunoplates on a levelling table.
Allow to set and air dry without a lid for 5 minutes.
iii)
Place lids on plates and store at 4°C in a humid box until used.
iv)
Prepare control plates with unsensitised cells or cells sensitised with an unrelated virus. Batches of
prepared plates can be stored for several weeks.
v)
Punch 3 mm holes in the set gels to a prepared template, allow for 16 test sera and a positive control
serum. On antigen control plates, prepare five rows of eight wells.
vi)
Pipette 10 µl of heat-inactivated (56°C for 30 minutes) test sera and a positive control serum to
appropriate wells. Incubate at 34°C for 20 hours in a humid box.
vii)
Measure zone diameters, and calculate areas of haemolysis after the area of the well has been
deducted.
•
Interpretation of the results
For results to be valid, positive and negative control sera should give results assigned through the
international collaborative study (Daly et al., 2007; Mumford, 2000) or if not using these international
standards, results consistent with those expected on the basis of prior experience. Areas of haemolysis for
the control sera should be clear and intra-laboratory variation should be no more than 5% for the control
serum. Results may be expressed as mm2 or as a ratio of the control serum value. Sera giving positive
results in the control plate should be adsorbed with sheep RBCs. For diagnostic purposes, acute and
convalescent sera should be tested in duplicate on the same plate. Increase in zone areas produced by
convalescent serum compared with acute serum is evidence of infection. The increase in area deemed to be
significant depends on the reproducibility of the test within the laboratory, but should be equivalent to a
twofold or more increase in antibody concentration. This area can be calculated from a standard curve
generated from a dilution series of a standard antiserum.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
1.
Background
a)
Rationale and intended use of the product
Equine influenza is a self-limiting disease and its economic significance is primarily due to the contagious
nature of the virus and the disruption to equestrian activities. Rapid diagnosis, movement restrictions and
vaccination are the key control measures for equine influenza. Vaccination decreases clinical signs and virus
2
Prepare three plates by adding 0.6, 1.2 or 1.8 ml of virus antigen to 2 ml RBCs. Add 1.3, 1.6 and 1.9 ml CrCl3 respectively
and resuspend to 2 ml in PBS ‘A’. Optimum volume of virus antigen is that which results in the largest and clearest zones
with appropriate reference serum.
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shedding. Influenza vaccines are widely available and are routinely used in competition horses in Europe,
the Americas, and Asia. In some countries, vaccination is mandatory for sport horses and racehorses that
are competing under rules of equestrian organisations. Following a primary course of three doses at
intervals of around 0, 1 and 6 months, an annual booster is the usual minimum requirement. More frequent
vaccination is recommended for young horses and some equestrian organisations require biannual
boosters.
Equine influenza virus vaccines typically consist of inactivated whole viruses or their glycoprotein subunits,
with or without adjuvant. Live attenuated virus and canary pox vectored vaccines have become available
commercially in some countries. Immunity generated by inactivated vaccines administered via the
intramuscular route is reliant on stimulation of circulating antibody to the HA, which neutralises virus; some
products have been shown to stimulate antibody in respiratory secretions. Critically the integrity and
conformation of the HA should be maintained during inactivation procedures to ensure that the vaccine
stimulates appropriate neutralising antibody. This can be tested by use of an immunological assay such as
SRD (single radial diffusion), which measures immunologically active HA capable of reacting with specific
anti-HA antibodies. The immunogenicity of the vaccine can be confirmed by measurement of HA antibody
stimulated in small animal models or the target species. An immune stimulating complex (ISCOM) based
vaccine against equine influenza has also been shown to stimulate systemic IFN gamma synthesis.
Canary pox-vectored vaccines, also administered via the intramuscular route, have the potential for cellmediated immunity (CMI) priming, as indicated, by increased equine influenza virus-specific IFN gamma and
IL-2 mRNA expression in vitro. Infectious titre of the recombinant canary pox virus carrying equine influenza
HA genes is used as an in vitro measure of potency and immunogenicity is assessed by measurement of
antibody stimulated in the target species.
Antibody to HA as measured by SRH, stimulated by inactivated whole virus, subunit or canary pox-vectored
vaccine correlates well with protection against infection in an experimental challenge model system (Edlund
Toulemonde et al., 2005; Mumford, 1983; 1994). In contrast, a cold-adapted temperature-sensitive mutant
used as a live attenuated vaccine replicates in the upper respiratory tract and does not stimulate high levels
of circulating antibody to HA but nevertheless provides protection against challenge infection. Immunity is
presumed to be mediated through mucosal or cellular responses rather than circulating antibody. As with the
vectored vaccine, in-process control testing is reliant on measurement of the infectious virus titre.
Guidelines for the production of veterinary vaccines are given in Chapter 1.1.6 Principles of veterinary
vaccine production. The guidelines given here and in chapter 1.1.6 are intended to be general in nature as
manufacturers are obliged to meet European Pharmacopeia, USDA or other national and regional
requirements.
All equine influenza vaccines should contain epidemiologically relevant strains
A formal equine influenza global surveillance programme has been in place since 1995.
The OIE Reference Laboratories and other collaborating laboratories collect data on
outbreaks of equine influenza and strain variation that is reviewed annually by an Expert
Surveillance Panel (ESP) including representatives from OIE and WHO. This panel
makes recommendations on the need to update vaccines, and these are published
annually in the OIE Bulletin (Expert Surveillance Panel, 2011). The criteria for updating
equine influenza vaccines are similar to those for human influenza vaccines and based
on analysis of evidence of disease in well vaccinated horses, antigenic changes, genetic
changes and, when possible, supporting experimental challenge data.
H7N7: Many vaccines still contain an H7N7 strain. However, the Expert Surveillance
Panel has recommended that the H7N7 component should be omitted as there have
been few reports of infections with this subtype have been substantiated during the past
30 years.
H3N8: Antigenic variants of H3N8 viruses co-circulate (Bryant et al., 2009) and it is
important to include a strain or strains that are epidemiologically relevant as
recommended by the OIE Expert Surveillance Panel and published in the OIE Bulletin
(Expert Surveillance Panel, 2010). OIE Reference Laboratories can provide help in
selecting suitable vaccine strains.
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Chapter 2.5.7. — Equine influenza
2.
Outline of production and minimum requirements for conventional vaccines
a)
Characteristics of the seed
i)
Biological characteristics
For each vaccine strain, a prototype batch should be prepared to establish its suitability as a vaccine
strain, i.e. purity and safety should be confirmed by standard techniques. The ability of seed-lot viruses
to grow to high titre and generate sufficient antigenic mass to stimulate adequate antibody responses
in the target species, should be confirmed. Additionally, vaccine virus derived in MDCK cells should be
fully characterised to ensure that antigenic variants have not arisen during the culture process, such
that the vaccine virus is no longer representative of the original isolate.
ii)
Quality criteria (sterility, purity, freedom from extraneous agents)
Virus strains may be obtained from OIE Reference Laboratories (see Table given in Part 4 of this
Terrestrial Manual). Viruses selected as vaccine strains should be described in terms of origin and
passage history. The strains are propagated in the allantoic cavity of 10-day-old embryonated hens’
eggs or cell cultures, such as MDCK. All manipulations must be conducted separately for each strain.
Viral growth is monitored by HA tests. Passaged virus is identified by serological tests, such as HI or
SRH or by PCR using specific primers. If vaccine virus is grown in cell culture, antigenic studies with
ferret sera and MAbs should be undertaken to ensure that variant viruses have not been selected
during passage to prepare master and working seed viruses. Master and working seed viruses are
divided into aliquots and stored in freeze-dried form at –20°C or at –70°C following testing for
extraneous agents. Records of storage conditions should be maintained.
The master seed lot of each vaccine strain selected should be processed at one time to assure a
uniform composition, tested for extraneous agents, and fully characterised. Antisera or MAbs for use in
HI tests to characterise vaccine strains may be obtained from OIE and WHO Reference Laboratories.
Working seed lots are derived from a master seed lot and should be of uniform composition, free from
extraneous agents, and fully characterised. Aliquots of the working seed are used for production of
vaccine.
Master and working seed lots should be prepared in specific pathogen free eggs or, as a minimum, in
eggs derived from a healthy flock.
If MDCK cells are used to propagate vaccine virus, master cell lines should be established and stored
in liquid nitrogen, and should be tested for freedom from extraneous agents according to National
Control Authority Guidelines.
Examination of seed viruses for extraneous agents including mycoplasmas and other equine viruses
should be performed by appropriate techniques, including inoculation of susceptible tissue cultures and
examination for cytopathic effect or application of fluorescent antibodies for antigen detection.
The presence of other common equine respiratory pathogens, e.g. equine herpesviruses 1, 2, 4,
equine picornaviruses, equine viral arteritis, and equine adenoviruses, should be specifically excluded.
The absence of bacteria should be confirmed by standard sterility tests and toxicity tests in small
animals.
b)
Method of manufacture
i)
Procedure
Production is based on a seed-lot system that has been validated with respect to the characteristics of
the vaccine strains. Where eggs are used, each strain of virus is inoculated separately into the allantoic
cavity of 9–11-day-old embryonated hens’ eggs from a healthy flock. The eggs are incubated at a
suitable temperature for 2–3 days, and the allantoic fluid is collected. Alternatively, each strain is
inoculated separately into MDCk cell cultures. The viral suspensions of each strain are collected
separately and inactivated. If necessary, they may be purified. Suitable adjuvants and antimicrobial
preservatives may be added.
Monovalent virus pools should be inactivated as soon as possible after their preparation, by a method
approved by the National Control Authority. If formalin (37% formaldehyde) or beta-propiolactone (2oxetanone) is used, the concentration by volume should not exceed 0.2%. Ideally, pools should be held
at 4°C and should be inactivated within 5 days of harvest. Inactivation of the vaccine must be
demonstrated. A suitable method consists of inoculating 0.2 ml of undiluted monovalent pool and 1/10
and 1/100 dilutions of the monovalent pool into the allantoic cavities of groups of fertile eggs (10 eggs
in each group), and incubating the eggs at 33–37°C for 3 days. At least 8 of the 10 eggs should survive
at each dosage level. A volume of 0.5 ml of allantoic fluid is harvested from each surviving egg. The
fluid harvested from each group is pooled, and 0.2 ml of each of the three pools is inoculated,
undiluted, into a further group of 10 fertile eggs. Haemagglutinin activity should not be detected in
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Chapter 2.5.7. — Equine influenza
these new groups of eggs. In some countries, the requirement that 80% of the eggs should survive
during incubation may be impossible to satisfy, in which case the National Control Authority should
then specify a modified requirement to be satisfied. Before inactivation, samples should be collected for
bacterial and fungal sterility tests.
Monovalent material may be concentrated and purified by high-speed centrifugation or other suitable
methods approved by the National Control Authority, either before or after the inactivation procedure. It
is important to concentrate and purify the virus under optimum conditions, e.g. temperatures that
preserve its antigenic properties.
The monovalent virus pool shall be shown not to contain viable influenza virus when tested by
inoculation of embryonated hens’ eggs, by a method approved by the National Control Authority.
Alternatively, the satisfactory inactivation can also be demonstrated by inoculating monolayers of
MDCK cells.
ii)
Requirements for substrates and media
Vaccines should be produced in eggs or in a cell line that meet the requirements of the National
Control Authority. Wherever practicable the use of substances of animal origin for example, serum and
trypsin should be kept to a minimum. Substances of animal origin used during production should be
subjected to a suitable, validated sterilisation or inactivation procedure or be tested for the absence of
extraneous organisms.
iii)
In-process controls
Relevant in-process controls should be applied before and after inactivation and before and after
concentration and purification.
In-process controls include: (a) identity of virus strains (tested by HI); (b) sterility; (c) virus titre;
(d) haemagglutinin content (tested by chicken RBCs agglutinating units, CCA [chick cell agglutination]);
and (e) immunologically active HA (tested by SRD or another suitable immunochemical method).
•
Single radial diffusion test
SRD is a reliable method for measuring immunologically active HA in terms of µg HA, and is used
routinely for potency testing of human influenza vaccines (Wood et al., 1983b).
The potency of inactivated equine influenza vaccine depends on the concentration of immunologically
active haemagglutinin (Wood et al., 1983a).
Assessment of the antigenic content of the vaccine by CCA alone may be misleading, as the sensitivity
of this assay is a reflection of the ability of virus strains to agglutinate RBCs. Disruption of virus may
lead to an apparent increase in HA as measured by CCA. The CCA assay does not provide a measure
of the antigenic properties of the HA (HA may retain its properties to bind to RBCs while losing its
ability to stimulate antibody).
Most equine influenza vaccines contain more than one variant of the H3N8 subtype. In this situation, it
is not possible to judge the potency of individual H3N8 components from serological tests performed
on sera collected from horses or small animals vaccinated with the final product, because of crossreactivity between the two isolates of the same subtype. Thus, it is important that a reliable method,
such as SRD, be used to measure the potency of individual components before and after inactivation
and prior to mixing and formulation with adjuvant.
In the SRD test, virus preparations are compared with a calibrated reference preparation of known HA
content. Antigens are allowed to diffuse through a gel containing an antiserum specific for a particular
HA. The distance diffused by the antigen before precipitation by the antibody incorporated in the gel is
directly related to the concentration of haemagglutinin in the antigen preparations.
Standard reagents for SRD testing are available from the WHO International Laboratory for Biological
Standards3. Reagents for A/eq/Prague/56 (H7N7), and the H3N8 strains A/eq/Miami/63,
A/eq/Kentucky/81, A/eq/Newmarket/1/93 (American lineage) and A/eq/Newmarket/2/93 (European
lineage) are currently available.
iv)
Final product batch tests
Sterility and purity
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
3
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National Institute for Biological Standards and Control, Blanche Lane, South Mimms, Potters Bar, Hertfordshire EN6 3QG,
UK.
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Chapter 2.5.7. — Equine influenza
Safety
a)
For inactivated or subunit vaccines absence of viral infectivity should be demonstrated by two
passages in 10 embryonated hen eggs. The allantoic fluid should have no haemagglutinating
activity.
b)
Using no fewer than three horses, each horse is inoculated intramuscularly (at two different sites)
with the dose of vaccine specified by the manufacturer; these inoculations are repeated 2–
4 weeks later. The animals are kept under observation for 10 days after the second set of
injections. No abnormal local or systemic reaction should ensue.
c)
If vaccine is to be used in mares, safety should be demonstrated by giving two doses of vaccine
to no fewer than two pregnant mares at the prescribed interval within the trimester for which the
vaccine is recommended. Once safety has been demonstrated on a prototype batch, safety
testing in pregnant mares may be omitted for routine testing of subsequent batches of the final
product.
Batch potency
Following mixing of viral antigens and adjuvants, aliquots should be potency tested in vivo using horses
and guinea-pigs or a suitable alternative immunochemical assay. Adjuvants cause interference in
quantitative in-vitro tests, such as CCA and SRD, although SRD may be used on the final product as a
qualitative assay to demonstrate the presence of antigen for each vaccine strain. For repeated batch
tests, only guinea-pigs or a suitable alternative immunochemical assay are used, subject to agreement
of the National Control Authority.
a)
Serological responses in horses
For a valid in-vivo potency test, naive seronegative horses must be selected for vaccination.
Young horses or ponies (not less than 6 months old) should be screened for the presence of
antibody using H7N7 and H3N8 viruses including recently isolated viruses relevant to the area in
which the horses were reared. If HI tests are used for screening, H3N8 viruses should be treated
with Tween 80/ether to maximise the sensitivity of the test. Alternatively, SRH may be used to
establish the seronegative status of animals.
To test a vaccine for efficacy in horses, inject a volume corresponding to one vaccine dose by the
recommended route into each of five susceptible seronegative horses. After the period
recommended between the first and second doses, as stated on the label, a volume of vaccine
corresponding to the second dose of vaccine is injected into each horse.
Three blood samples are collected from each animal, the first at the time of the first vaccination,
the second 1 week after the first vaccination to check for anamnestic response, and the third
2 weeks after the second vaccination.
The serological assay used to measure the antibody response to the viruses contained in the
vaccine must be standardised for a valid in-vivo potency test. The SRH assay (see Section B.2.b)
is preferred as standard reference sera are available for quality control purposes from the
European Pharmacopoeia4. These sera should be tested in parallel with the test sera to ensure
that the test is valid with respect to sensitivity; the values obtained should not vary by more than
20% from the SRH values assigned in an international collaborative study (Daly et al., 2007;
Mumford, 2000). Due to poor repeatability and reproducibility of the HI test between different
laboratories, no HI titre could be assigned to these sera.
Following vaccination the antibody value measured by SRH should not be less than 150 mm2.
This is higher than the value required in the European Pharmacopoeia Monograph for inactivated
equine influenza vaccines (85 mm2) as this value is not considered to be protective. If the value
found for any horse after the first vaccination indicates that there has been an anamnestic
response, the result is not taken into account. A supplementary test is carried out, as described
above, replacing the horses that showed an anamnestic response with an equal number of new
animals.
If the HI test is used, the antibody titre of each serum taken after the second vaccination in each
test should not be less than 1/64 (calculated for the original serum, taking into account the
predilution of 1/8). Alternatively, the antibody levels stimulated by the vaccine under test should
be shown to be at least equal to the antibody levels stimulated by a standard vaccine tested in
parallel that has been shown previously to protect horses against challenge infection.
4
Serum to A/eq/Newmarket/1/77
A/eq/Newmarket/2/93 (E0850022).
OIE Terrestrial Manual 2012
(Catalogue
number
E0850010),
A/eq/Newmarket/1/93
(E0850021)
and
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Chapter 2.5.7. — Equine influenza
b)
Challenge studies in horses
It may be desirable in certain cases to undertake challenge studies in horses to demonstrate
potency, particularly if vaccines are being assessed for their ability to protect against antigenically
dissimilar viruses. Challenge studies may be carried out by exposing vaccinated horses/ponies to
an aerosol of virulent influenza virus no fewer than 2 weeks after the second dose of vaccine.
Comparisons of clinical signs, virus excretion and serological responses are made with a group of
unvaccinated control animals challenged at the same time. The timing of the challenge procedure
will reflect the claims to be made on the data sheet regarding duration of immunity. Protection at
2 weeks post-vaccination when antibody levels are at their peak level does not necessarily
indicate good duration of immunity under field conditions.
If tests for potency in horses have been carried out with satisfactory results on a representative
batch of vaccine, these tests may be omitted as a routine control on other batches of vaccine
prepared using the same seed-lot system, subject to agreement by the National Control Authority.
c)
Serological responses of guinea-pigs
Inject each of no fewer than five guinea-pigs free from specific antibodies with one vaccine dose.
Collect blood samples 21 days later, and test the serum by SRH or HI (see Sections B.2.a and
B.2.b). Perform the tests of each serum using, respectively, the antigen(s) prepared from the
strain(s) used in the production of the vaccine. The antibody titre of each serum in each test
should not be less than the titre stimulated by a standard vaccine that has been shown to
stimulate protective levels of antibody in horses.
c)
Requirements for authorisation
i)
Safety requirements
The safety requirements may vary with the National Authority but usually include the assessment of the
administration of an overdose and of the repeated administration of one dose to young horses and
pregnant mares, if the vaccine is intended for use in pregnant mares. For live vaccines the
dissemination by the vaccinated horse and the spread of the vaccine strain from vaccinated to
unvaccinated horses along with the possible consequences must be investigated. Reversion to
virulence studies should be performed by serial passaging of the live vaccine.
ii)
Efficacy requirements
The efficacy requirements may vary with the National Authority but usually include the assessment of
the serological response in horses and virus challenge studies in susceptible horses (see Section
C.2.b.iv batch potency b).
Where claims for duration of immunity are made on the data sheet, these should be supported with
data on the duration of protective levels of antibody maintained in horses vaccinated according to the
recommended schedule. Antibody levels quoted as protective should be validated in challenge studies
or by comparison with published reports.
iii)
Stability
Vaccines should be stored at 5±3°C and protected from light. The shelf life quoted on the data sheet
should be demonstrated by testing the potency of aliquots over time using the guinea-pig potency test
(see Section C.2.b.iv batch potency c).
iv)
Maintaining epidemiologically relevant strains in vaccines
To enable vaccine manufacturers to respond quickly to recommendations from the Expert Surveillance
Panel to update vaccine strains, the Committee for Veterinary Medicinal Products (CVMP) for the
European Agency for the Evaluation of Medicinal Products (EMEA) has developed a fast-track
licensing system to be used when vaccine strains are updated (EMEA, 1998).
3.
Vaccines based on biotechnology
a)
Vaccines available and their advantages
A canary pox recombinant vaccine is available in some countries and was used in the equine influenza
control and eradication programme in Australia in 2007. A nucleoprotein ELISA was used to differentiate
horses vaccinated with the recombinant vaccine from horses that had been exposed to virus by natural
infection (DIVA) This DIVA was possible because the canary pox recombinant expresses only the
haemagglutinin gene of equine influenza. There is some evidence that this vaccine may be used to prime
the immune system in the face of maternally derived antibodies (Minke et al., 2008).
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Chapter 2.5.7. — Equine influenza
b)
Special requirements
National Authorities frequently require an environmental risk assessment. This may include an assessment
of the potential direct and indirect, immediate or delayed adverse effects of the genetically modified vaccine
on human and animal health and on the environment. The phenotypic and genetic stability of the vaccine
and its potential interactions with other organisms may need to be evaluated.
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JOHN T.J. & FULGINITI V.A. (1966). Parainfluenza 2 virus: increase in haemagglutinin titre on treatment with Tween80 and ether. Proc. Soc. Exp. Biol. Med., 121, 109–111.
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LIVESAY G.J., O’NEILL T., HANNANT D, YADAV M.P. & MUMFORD J.A. (1993). The outbreak of equine influenza (H3N8)
in the United Kingdom in 1989; diagnostic use of an antigen capture ELISA. Vet. Rec., 133, 515–519.
LU Z., CHAMBERS T.M., BOLIAR S., BRANSCUM A.J., STURGILL T.L., TIMONEY P.J., REEDY S.E., TUDOR L.R., DUBOVI
E.J., VICKERS M.L., SELLS S. & BALASURIYA U.B. (2009). Development and evaluation of one-step Taqman real-time
reverse transcription-PCR assays targeting nucleoprotein, matrix and hemagglutinin genes of equine influenza
virus. J. Clin. Microbiol., 47, 3907–3913.
MINKE J.M., TOULEMONDE C.E., DINIC S., COZETTE V, CULLINANE A., AUDONNET J.C. (2007). Effective priming of foals
born to immune dams against influenza by a canarypox-vectored recombinant influenza H3N8 vaccine. J. Comp.
Pathol., 137, S76–S80.
MORLEY P.S., BOGDAN J.R., TOWNSEND H.G.G. & HAINES D.M. (1995). The effect of changing single radial
haemolysis assay method when quantifying influenza A antibodies in serum. Vet. Microbiol., 44, 101–110.
MUMFORD J.A. (1992). Progress in the control of equine influenza. In: Equine Infectious Disease VI: Proceedings
of the Sixth International Conference, Plowright W., Rossdale P.D. & Wade J.F., eds. Newmarket, R & W
Publications, UK, 207–217.
MUMFORD J. (2000). Collaborative study for the establishment of three European Pharmacopoeia biological
reference preparations for equine influenza horse antiserum. PHARMEUROPA Special Issue, Bio 2000-1, 5–21.
MUMFORD J.A., JESSETT D., DUNLEAVY U., WOOD J., HANNANT D., SUNDQUIST B. & COOK R.F. (1994). Antigenicity and
immunogenicity of experimental equine influenza ISCOM vaccines. Vaccine, 12, 857–863.
MUMFORD J., WOOD J.M., SCOTT A.M., FOLKERS C. & SCHILD G.C. (1983). Studies with inactivated equine influenza
vaccine 2. Protection against experimental infection with influenza virus A/equine/Newmarket/79 (H3N8). J. Hyg.
(Camb.), 90, 385–395.
OXBURGH L. & KLINGBORN B. (1999). Cocirculation of two distinct lineages of equine influenza virus subtype H3N8.
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QUINLIVAN M., DEMPSEY E., RYAN F., ARKINS S. & CULLINANE A. (2005). Real-time reverse transcription PCR for
detection and quantitative analysis of equine influenza virus. J. Clin. Microbiol., 43, 5055–5057.
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characterization of equine H3N8 influenza virus from pigs in China. Arch. Virol., 154, 887–890.
WEBSTER R.G. (1993). Are equine 1 influenza viruses still present in horses? Equine Vet. J., 25, 537–538.
WOOD J.M., MUMFORD J., FOLKERS C., SCOTT A.M. & SCHILD G.C. (1983b). Studies with inactivated equine influenza
vaccine 1. Serological responses of ponies to graded doses of vaccine. J. Hyg. (Camb.), 90, 371–384.
WOOD J.M., SCHILD G.C., FOLKERS C., MUMFORD J. & NEWMAN R.W. (1983a). The standardisation of inactivated
equine influenza vaccines by single-radial immunodiffusion. J. Biol. Stand., 11, 133–136.
YAMANAKA T., TSUJIMURA K., KONDO T. & MATSUMURA T. (2008). Evaluation of antigen detection kits for diagnosis of
equine influenza. J. Vet. Med. Sci., 70, 189–192.
*
* *
NB: There are OIE Reference Laboratories for Equine influenza
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for equine influenza
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.8.
EQUINE PIROPLASMOSIS
SUMMARY
Equine piroplasmosis is a tick-borne protozoal disease of horses, mules, donkeys and zebra. The
aetiological agents are blood parasites named Theileria equi and Babesia caballi. Theileria equi
was previously designated as Babesia equi. Infected animals may remain carriers of these
parasites for long periods and act as sources of infection for ticks, which act as vectors. The
parasites are found inside the red blood cells of the infected animals.
The introduction of carrier animals into areas where tick vectors are prevalent can lead to an
epizootic spread of the disease.
Identification of the agent: Infected horses can be identified by demonstrating the parasites in
stained blood or organ smears during the acute phase of the disease. Romanovsky-type staining
methods, such as Giemsa, give the best results. In carrier animals, low parasitaemias make it
extremely difficult to detect parasites, especially in the case of B. caballi infections, although they
may sometimes be demonstrated by using a thick blood smear technique.
Paired merozoites joined at their posterior ends are a diagnostic feature of B. caballi infection. The
parasites in the erythrocytes measure 2 × 5 µm. The merozoites of T. equi are less than 2–3 µm
long, and are pyriform, round or ovoid. A characteristic of T. equi is the arrangement of four pearshaped merozoites forming a tetrad known as a ‘Maltese cross’.
Molecular techniques for the detection of T. equi and B. caballi based on species-specific
polymerase chain reaction (PCR) assays, targeting the 18S rRNA gene, have been developed and
continue to expand. These tests have been shown to be highly specific and sensitive and promise
to play an increasing role in the diagnosis of infections.
Serological tests: Infections in carrier animals are best demonstrated by testing their sera for the
presence of specific antibodies.
Currently, the indirect fluorescent antibody (IFA) test and the competitive enzyme-linked
immunosorbent assay (C-ELISA) are the primary tests used for qualifying horses for importation.
The complement fixation (CF) test, for many years the primary test, has been replaced by the IFA
and C-ELISA. These tests have proven to be more effective at detecting long-term infected animals
and animals treated with antiparasitic drugs; these animals may be CF negative but still be infected.
The IFA test and C-ELISA have been shown to be highly specific for each of the two species of
piroplasmosis agents involved. Indirect ELISAs may also be used to detect antibodies to both
species in infected horses, although cross-reactions between T. equi and B. caballi occur.
Application of recombinant T. equi and B. caballi merozoite proteins in diagnostic assays appear to
be very promising in the accurate determination of equine piroplasmosis infection.
Requirements for vaccines and diagnostic biologicals: There are no biological products
available.
A. INTRODUCTION
Equine piroplasmosis is a tick-borne protozoal disease of horses, mules, donkeys and zebra. The aetiological
agents of equine piroplasmosis are Theileria equi and Babesia caballi. Twelve species of Ixodid ticks in the
genera Dermacentor, Rhipicephalus and Hyalomma have been identified as transstadial vectors of B. caballi and
T. equi, while eight of these species were also able to transmit B. caballi infections transovarially (De Waal &
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Chapter 2.5.8. — Equine piroplasmosis
Potgieter, 1987; Neitz, 1956; Stiller & Frerich, 1979; Stiller et al., 1980). Infected animals may remain carriers of
these blood parasites for long periods and act as sources of infection for tick vectors.
The parasites occur in southern Europe, Asia, countries of the Commonwealth of Independent States, Africa,
Cuba, South and Central America, and certain parts of the southern United States of America. Theileria equi has
also been reported from Australia (but, apparently never established itself in this region), and is now believed to
have a wider general distribution than B. caballi.
During the life cycle of Babesia, initially the sporozoites invade red blood cells (RBCs) where they transform into
trophozoites (Friedhoff, 1970; Potgieter & Els, 1977). In this situation the trophozoites grow and divide into two
round, oval or pear-shaped merozoites. The mature merozoites are now capable of infecting new RBCs and the
division process is then repeated.
Babesia caballi: the merozoites in the RBCs are pear-shaped, 2–5 µm long and 1.3–3.0 µm in diameter (Levine,
1985). The paired merozoites joined at their posterior ends are considered to be a diagnostic feature of B. caballi
infection (Mehlhorn & Schein, 1984).
Theileria equi: the merozoites of this organism are relatively small, less than 2–3 µm long (Levine, 1985), and are
pyriform, round or ovoid. A characteristic of T. equi is the arrangement of four pear-shaped merozoites,
measuring about 2 µm in length, forming a tetrad known as the ‘Maltese cross’ arrangement (Holbrook et al.,
1968).
In T. equi infection it has been shown that sporozoites inoculated into horses via a tick bite invade the
lymphocytes (Schein et al., 1981). The sporozoites undergo development in the cytoplasm of these lymphocytes
and eventually form Theileria-like schizonts. Merozoites released from these schizonts enter RBCs. The
taxonomic position of T. equi has been controversial and only relatively recently has it been redescribed as a
Theileria (Mehlhorn & Schein, 1998). Further support for the close relation with Theileria spp. also comes from the
homology found between 30 and 34 kDa T. equi surface proteins and similar sized proteins of various Theileria
spp. (Kappmeyer et al., 1993; Knowles et al., 1997). However, the position of T. equi in phylogenetic trees based
on the small subunit ribosomal RNA genes is variable and mostly appear as a sister clade of the Theilerids
(Criado-Fornelio et al., 2003; Nagore et al., 2004) leading some to suggest that T. equi is ancestral to the
Theilerids (Criado-Fornelio et al., 2003) or a different group altogether (Allsopp et al., 1994).The clinical signs of
equine piroplasmosis are often nonspecific, and the disease can easily be confused with other conditions.
Piroplasmosis can occur in peracute, acute and chronic forms. The acute cases are more common, and are
characterised by fever that usually exceeds 40°C, reduced appetite and malaise, elevated respiratory and pulse
rates, congestion of mucous membranes, and faecal balls that are smaller and drier than normal.
Clinical signs in subacute cases are similar. In addition, affected animals show loss of weight, and fever is
sometimes intermittent. The mucous membranes vary from pale pink to pink, or pale yellow to bright yellow.
Petechiae and/or ecchymoses may also be visible on the mucous membranes. Normal bowel movements may be
slightly depressed and the animals may show signs of mild colic. Mild oedematous swelling of the distal part of
the limbs sometimes occurs.
Chronic cases usually present nonspecific clinical signs such as mild inappetence, poor performance and a drop
in body mass. The spleen is usually found to be enlarged on rectal examination.
A rare peracute form where horses are found either dead or moribund has been reported (Littlejohn, 1963).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
Infected horses may be identified by demonstrating the parasites in stained blood, optimally collected from
superficial skin capillaries, or organ smears during the acute phase of the disease. Romanovsky-type staining
methods, such as the Giemsa method, usually give the best results (Shute, 1966). However, even in acute clinical
cases of B. caballi infection, the parasitaemia is very low and difficult to detect. Experienced workers sometimes
use a thick blood smear technique (Mahoney & Saal, 1961) to detect very low parasitaemia. Thick films are made
by placing a small drop (approximately 50 µl) of blood on to a clean glass slide which is then air-dried, heat fixed
at 80°C for 5 minutes, and stained in 5% Giemsa for 20–30 minutes.
An accurate identification of the species of the parasite is sometimes desirable, as mixed infections of T. equi and
B. caballi probably occur frequently.
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Chapter 2.5.8. — Equine piroplasmosis
Identification of equine piroplasmosis in carrier animals by means of blood smear examination is not only very
difficult but also inaccurate and therefore serological methods are preferred for this (see below). However, falsenegative or false-positive reactions may be encountered in the course of serological tests (Donnelly et al., 1980;
Frerichs et al., 1969; Tenter & Freidhoff, 1986). In such cases, the passage of whole blood, although a
cumbersome and expensive exercise, is a very useful technique to determine the true position. Large quantities of
whole blood (500 ml) are transfused into a susceptible, preferably splenectomised, horse. This animal is then kept
under close observation for clinical signs of disease. Diagnosis is confirmed by the presence of parasites in its
RBCs.
In an additional technique, a specific uninfected tick vector is fed on a suspect animal, and the organism can then
either be identified in the tick itself or through the transmission of the organism by the tick vector to another
susceptible animal.
Success in the establishment of in-vitro cultures of T. equi and B. caballi may be one alternative to supplement
the methods described above, in order to identify carriers of the parasites (Holman et al., 1993; 1994; Zweygarth
et al., 1995; 1997). Babesia caballi parasites were successfully cultured from the blood of two horses that tested
negative by the complement fixation (CF) test (Holman et al., 1993). Similarly, T. equi could be cultured from
horses that did not show any patent parasitaemia at the time of the initiation of the cultures (Zweygarth et al.,
1995; 1997).
Molecular techniques for the detection of T. equi and B. caballi have been described. These methods are based
on species-specific polymerase chain reaction (PCR) assays, which mainly target the 18S rRNA gene
(Bashiruddin et al., 1999; Criado-Fornelio et al., 2003; Sahagun-Ruiz et al., 1997). Further refinements to the
technique includes nested PCR (Rampersad et al., 2003), loop-mediated isothermal amplification (LAMP)
(Alhassan et al., 2007) with reported increased sensitivity; a highly sensitive reverse line blot assay (RLB) and
multiplex PCR for simultaneous detection and identification of Theileria and Babesia species in horses (Alhassan
et al., 2005; Nagore et al., 2004).
2.
Serological tests
It is extremely difficult to diagnose the organisms in carrier animals by means of the microscopic examination of
blood smears. Furthermore, it is by no means practical on a large scale. The serological testing of animals is
therefore recommended as a preferred method of diagnosis, especially when horses are destined to be imported
into countries where the disease does not occur, but the vector is present.
Sera should be collected and dispatched to diagnostic laboratories in accordance with the specifications of that
laboratory. Horses for export that have been subjected to serological tests and shown to be free from infection,
should be kept free of ticks to prevent accidental infections.
A number of serological techniques have been used in the diagnosis of piroplasmosis, such as the CF test, the
indirect fluorescent antibody (IFA) test and the enzyme-linked immunosorbent assay (ELISA). In addition, a
simple and rapid immunochromatographic test for T. equi has also recently been described and might be a very
useful test for the mass screening of serum samples (Huang et al., 2004).
a)
Indirect fluorescent antibody test (a prescribed test for international trade)
The IFA test has been successfully applied to the differential diagnosis of T. equi and B. caballi infections
(Madden & Holbrook, 1968). The recognition of a strong positive reaction is relatively simple, but any
differentiation between weak positive and negative reactions requires considerable experience in
interpretation. A detailed description of the protocol of the IFA test has been given (Madden & Holbrook,
1968; Morzaria et al., 1977). An example of an IFA protocol is given below.

Antigen production
Blood for antigen is obtained from horses with a rising parasitaemia, ideally 2–5%. Carrier animals that have
already produced antibodies are not suitable for antigen production. Blood (about 15 ml) is collected into
235 ml of phosphate buffered saline (PBS), pH 7.2. The RBCs are washed three times in cold PBS (1000 g
for 10 minutes at 4°C). The supernatant fluid and the white cell layer are removed after each wash. After the
last wash, the packed RBCs are reconstituted to the initial volume with 4% bovine serum albumin fraction V
made up in PBS, i.e. the original packed cell volume = 30%, so that one-third consists of RBCs. If the
original RBC volume is 15 ml, then 5 ml of packed RBCs + 10 ml of 4% bovine albumin in PBS constitutes
the antigen. After thorough mixing, the antigen is placed on to prepared wells on a glass slide using a
template or a syringe (Morzaria et al., 1977). Alternatively, the cells can be spread smoothly on to
microscope slides, covering the entire slide with an even, moderately thick film. These slides are allowed to
dry, wrapped in soft paper and sealed in plastic bags or wrapped in aluminium foil, and stored at –20°C for
up to 1 year.
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
Test procedure
i)
Each sample of serum is tested against an antigen of B. caballi and of T. equi.
ii)
Prior to use, the frozen antigen slides are removed from storage at –20°C and incubated at 37°C for
10 minutes.
iii)
The antigen smears are then removed from their protective covering and fixed in cold dry acetone
(–20°C) for 1 minute. Commercially produced slides are available that are pre-fixed.
iv)
If smears were prepared on the whole slide surface, squares (14–21 in number, i.e. 2–3 rows of
7 each) are formed on the antigen smears with nail varnish or rapidly drying mounting medium (i.e.
Cystoseal).
v)
Test, positive and negative control sera are diluted from 1/80 to 1/1280 in PBS. Negative and positive
control sera are included in each test.
vi)
Sera are applied (10 µl each) at appropriate dilutions to the different wells or squares on the antigen
smear, incubated at 37°C for 30 minutes, and washed several times in PBS and once in water.
vii)
An anti-horse immunoglobulin prepared in rabbits and conjugated with fluorescein isothiocyanate (this
conjugate is available commercially) is diluted in PBS and applied to the smear, which is then
incubated and washed as before.
viii) After the final wash, two drops of a solution containing equal parts of glycerin and PBS are placed on
each smears and mounted with a cover-slip.
ix)
b)
The smear is then examined under the microscope for the fluorescing parasites. Sera diluted 1/80 or
more that show strong fluorescence are usually considered to be positive, although due consideration
is also given to the patterns of fluorescence of the positive and negative controls.
Enzyme-linked immunosorbent assay (a prescribed test for international trade)
In recent years a number of recombinant antigens for the use in ELISAs have been described. Recombinant
T. equi (EMA-1; EMA-2; Be82 and Be158) and B. caballi proteins (RAP-1; Bc48; Bc134) have been
produced in Escherichia coli (Hirata et al., 2002; 2005; Huang et al., 2003; Ikadai et al., 2000; Kappmeyer et
al., 1999; Knowles et al., 1992; Tamaki et al., 2004) or in insect cells by baculovirus (Xuan et al., 2001;
Tanaka et al., 1999). Recombinant antigens produced in E. coli or by baculovirus have the obvious
advantage of avoiding the need to infect horses for antigen production, and of eliminating the crossreactions that have been experienced in the past with the crude ELISA antigens. They also provide a
consistent source of antigen for international distribution and standardisation.
Indirect ELISAs using EMA-2 and BC48 have shown high sensitivity and specificity in detecting antibodies in
infected horses (Huang et al., 2003; Ikadai et al., 1999; Tanaka et al., 1999). Initial results from these tests
are promising and further validation of the assays is underway.
A competitive inhibition ELISA (C-ELISA) using EMA-1 protein and a specific monoclonal antibody (MAb)
that defines this merozoite surface protein epitope, have been used in a C-ELISA for T. equi (Knowles et al.,
1992). This C-ELISA overcomes the problem of antigen purity, as the specificity of this assay depends only
on the specificity defined by the MAb T. equi epitope. A 94% correlation was shown between the C-ELISA
and the CF test in detecting antibodies to T. equi. Sera that gave discrepant results were evaluated for their
ability to immunoprecipitate 35S-methionine-labelled in-vitro translated products of T. equi merozoite mRNA.
Samples that were C-ELISA positive and CF test negative clearly precipitated multiple T. equi proteins.
However, immunoprecipitation results with serum samples that were C-ELISA negative and CF test positive
were inconclusive (Knowles et al., 1991). This C-ELISA for T. equi was also recently validated in Morocco
and Israel, giving a concordance of 91% and 95.7% with the IFA test, respectively (Rhalem et al., 2001;
Shkap et al., 1998). A similar C-ELISA has been developed using the recombinant B. caballi rhoptryassociated protein 1 (RAP-1) and an MAb reactive with a peptide epitope of a 60 kDa B. caballi antigen
(Kappmeyer et al., 1999). The results of 302 serum samples tested with this C-ELISA and the CF test
showed a 73% concordance. Of the 72 samples that were CF test negative and C-ELISA positive, 48 (67%)
were shown to be positive on the IFA test, while four of the five samples that tested CF test positive and CELISA negative were positive on the IFA test (Kappmeyer et al., 1999).
A test protocol for an equine piroplasmosis C-ELISA has been described and used for additional validation
studies (Katz et al., 2000; United States Department of Agriculture [USDA], 2005). The apparent specificity
of the T. equi and B. caballi C-ELISAs lay between 99.2% and 99.5% using sera from 1000 horses
presumed to be piroplasmosis-free. One thousand foreign-origin horses of unknown infection status were
tested by the C-ELISA and the CF test with an apparent greater sensitivity of the C-ELISA. The results were
1.1% (T. equi) and 1.3% (B. caballi) more seropositive animals detected by C-ELISA than by the CF test; the
additional positive results were confirmed by IFA testing. A similar study of 645 foreign-origin horses tested
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Chapter 2.5.8. — Equine piroplasmosis
for import and pre-import purposes used heat-treated sera (58°C for 30 minutes), and resulted in 3.6%
(T. equi) and 2.1% (B. caballi) more seropositive animals detected by the C-ELISA than by the CF test. Both
C-ELISAs were highly reproducible well-to-well, plate-to-plate, and day-to-day, with overall variances of ±
1.2% and ±1.6% for the T. equi and B. caballi tests, respectively.
The C-ELISA protocol is given below.
A detailed description of antigen production and a test protocol has been given by the National Veterinary
Services Laboratories of the USDA (2005). A commercial kit is now available that is based on the same
antigens and monoclonal antibodies).

Solutions
Antigen coating buffer: prepare the volume of antigen coating buffer required using the following amounts of
ingredients per litre: 2.93 g sodium bicarbonate; 1.59 g sodium carbonate; sufficient ultra-pure water to
dissolve, and make up to 1 litre with ultra-pure water. Adjust to pH 9.6.
C-ELISA wash solution (high salt diluent): prepare the volume of C-ELISA wash solution required by using
the following amounts of ingredients per litre: 29.5 g sodium chloride; 0.22 g monobasic sodium phosphate;
1.19 g dibasic sodium phosphate; 2.0 ml Tween 20; sufficient ultra-pure water to dissolve, and make up to
1 litre with ultra-pure water. Mix well. Adjust pH to 7.4. Sterilise by autoclaving at 121°C.

Antigen production
Frozen transformed E. coli culture is inoculated at a 1/10,000 dilution into any standard non-selective
bacterial growth broth (e.g. Luria broth) containing added carbenicillin (100 µg/ml) and isopropylthiogalactoside (IPTG, 1 mM). Cultures are incubated on an orbital shaker set at 200 rpm at 37°C overnight.
Cells grown overnight are harvested by centrifugation (5000 g for 10 minutes), washed in 50 mM Tris/HCl
and 5 mM ethylene diamine tetra-acetic acid (EDTA) buffer, pH 8.0, and harvested again as before. (Antigen
is available from the National Veterinary Services Laboratories, P.O. Box 844, Ames, Iowa 50010, USA.)
Cells are resuspended to 10% of the original volume in the Tris/EDTA buffer to which 1 mg/ml of lysozyme
has been added, and incubated on ice for 20 minutes. Nonidet P-40 detergent (NP-40) is then added to a
final 1% concentration (v/v), vortexed, and the mixture is incubated on ice for 10 minutes. The material is
next sonicated four times for 30 seconds each time at 100 watts, on ice, allowing 2 minutes between
sonications for the material to remain cool. The sonicate is centrifuged at 10,000 g for 20 minutes. The
resulting supernatant is dispensed in 0.5 ml aliquots in microcentrifuge tubes and may then be stored at
–70°C for several years. The presence of heterologous host bacterial antigens does not interfere with the
binding of specific equine anti-piroplasma antibodies or the binding of the paired MAbs to their respective
expressed recombinant antigen epitopes and is confirmed by the following procedures. The antigencontaining supernatants are quality controlled by titrating them with their paired MAbs and with reference
monospecific equine antisera to verify both an adequate level of expression and complete specificity for the
homologous species of piroplasmosis agent. Normal serum (negative serum) controls must not interfere with
binding of the MAbs or positive equine reference sera to the expressed antigen preparation.

Test procedure
i)
Microtitration plates are prepared by coating the wells with 50 µl of either T. equi antigen or B. caballi
antigen diluted in antigen-coating buffer. The dilution used is determined by standard serological
titration techniques. The plate is sealed with sealing tape, stored overnight at 4°C, and frozen at
–70°C. Plates can be stored at –70 °C for up to 6 months.
ii)
The primary anti-T. equi or anti-B. caballi MAb and secondary antibody-peroxidase conjugate is diluted
as directed by the manufacturer at the time of use in the C-ELISA, with antibody-diluting buffer
(supplied with the test kit).
iii)
Plates are thawed at room temperature, the coating solution is decanted, and the plates are washed
twice with C-ELISA wash solution.
iv)
The serum controls and test serum samples are diluted 1/2 with serum-diluting buffer before 50 µl of
sera is added to wells. Each unknown serum sample is tested in single or duplicate wells. Positive
control sera and blanks are tested in duplicate while negative controls are tested in triplicate on
different parts of the plate. Plates are incubated covered, at room temperature (21–25°C) for
30 minutes in a humid chamber, and then washed three times in C-ELISA wash solution.
v)
All wells then receive 50 µl/well of diluted primary anti-T. equi or anti-B. caballi MAb. (The MAb is
produced in a cell culture bioreactor and is available from the National Veterinary Services
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Chapter 2.5.8. — Equine piroplasmosis
Laboratories, P.O. Box 844, Ames, Iowa 50010, USA.) Plates are incubated covered for 30 minutes at
room temperature (21–25°C) in a humid chamber, and then washed three times in C-ELISA wash
solution.
vi)
Diluted secondary peroxidase anti-murine IgG (50 µl/well) conjugate is added to each well. Plates are
incubated covered for 30 minutes at room temperature (21–25°C) in a humid chamber, and then
washed three times in C-ELISA wash solution.
vii)
Chromogenic enzyme substrate (50 µl/well) is added to all wells, and plates are incubated for
15 minutes at room temperature (21–25°C) during colour development.
viii) The colour development is stopped by adding 50 µl of stop solution to all wells and the plates are read
immediately on a plate reader.
c)
ix)
The plates are read at 620, 630 or 650 nm wavelength (OD). The average OD is calculated for the
duplicate wells for all control sera and blank wells. For a valid test, the mean of the negative controls
must produce an OD >0.300 and <2.000. The mean positive control sera must produce an inhibition of
≥40%.
x)
Per cent inhibition [%I] is calculated as follows: %I = 100 – [(Sample OD × 100) ÷ (Mean negative
control OD)].
xi)
If a test samples produces ≥40% inhibition it is considered positive. If the test sample produces <40%
inhibition it is considered negative.
Complement fixation
The CF test has been used in the past by some countries to qualify horses for importation (Taylor et al.,
1969). A detailed description of antigen production and a test protocol has been given (for example: USDA,
1997). The CF test may not identify all infected animals, especially those that have been drug-treated or that
produce anti-complementary reactions, or because of the inability of IgG(T) (the major immunoglobulin
isotype of equids) to fix guinea-pig complement (McGuire et al., 1971). Therefore the IFA test and C-ELISA
have replaced the CF as the prescribed tests for international trade.
An example of a CF test protocol is given below.

Solutions
Alsever’s solution: prepare 1 litre of Alsever’s solution by dissolving 20.5 g glucose; 8.0 g sodium citrate;
4.2 g sodium chloride in sufficient distilled water. Adjust to pH 6.1 using citric acid, and make up the volume
to 1 litre with distilled water. Sterilise by filtration.
Stock veronal buffer (5×): dissolve the following in 1 litre of distilled water: 85.0 g sodium chloride; 3.75 g
sodium 5,5 diethyl barbituric; 1.68 g magnesium chloride (MgCl2.6H2O); 0.28 g calcium chloride. Dissolve
5.75 g of 5,5 diethyl barbituric acid in 0.5 litre hot (near boiling) distilled water. Cool this acid solution and
add to the salt solution. Make up to 2 litres with distilled water and store at 4°C. To prepare a working
dilution, add one part stock solution to four parts distilled water. The final pH should be from 7.4 to 7.6.

Antigen production
Blood is obtained from horses with a high parasitaemia (e.g. 3–7% parasitaemia for B. caballi and 60–85%
for T. equi), and mixed with equal volumes of Alsever’s solution as an anticoagulant. The plasma/Alsever’s
supernatant and buffy coat are removed when the RBCs have settled to the bottom of the flask. The RBCs
are washed several times with cold veronal buffer and then disrupted. The antigen is recovered from the
lysate by centrifugation at 30,900 g for 30 minutes.
The recovered antigen is washed several times in cold veronal buffer by centrifugation at 20,000 g for
15 minutes. Polyvinyl pyrrolidone 40,000 (1–5%[ w/v]) is added as a stabiliser and the preparation is mixed
on a magnetic stirrer for 30 minutes, strained through two thicknesses of sterile gauze, dispensed into 2 ml
volumes and freeze-dried. The antigen can then be stored at below –50°C for several years.
884

Test procedure
i)
The specificity and potency of each batch of antigen should be checked against standard antisera of
known specificity and potency. Optimal antigen dilutions are also determined in a preliminary
checkerboard titration.
ii)
Test sera are inactivated for 30 minutes at 58°C (donkey and mule sera are inactivated at 62.5°C for
35 minutes) and tested in dilutions of 1/5 to 1/5120. Veronal buffer is used for all dilutions.
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Chapter 2.5.8. — Equine piroplasmosis
iii)
Complement is prepared and titrated spectrophotometrically to determine the 50% haemolytic dose
(C’H50) (Stiller et al., 1980) and used in the test at five times C’H50. The haemolytic system consists of
equal parts of a 2% sheep (RBC) suspension and veronal buffer with 5 minimum haemolytic doses
(MHDs) of haemolysin (amboceptor) (USDA, 1997). Some laboratories use twice the 100% haemolytic
dose, which gives equivalent sensitivity.
iv)
The test has been adapted to microtitration plates (Herr et al., 1985). The total volume of the test is
0.125 ml, made up of equal portions (0.025 ml) of antigen, complement (five times C’H50) and diluted
serum. Incubation is performed for 1 hour at 37°C.
v)
A double portion (0.05 ml) of the haemolytic system is added and the plates are incubated for a further
45 minutes at 37°C with shaking after 20 minutes.
vi)
The plates are centrifuged for 1 minute at 200 g before being read over a mirror.
vii)
A lysis of 50% is recorded as positive, with the titre being the greatest serum dilution giving 50% lysis.
A titre of 1/5 is regarded as positive. A full set of controls (positive and negative sera) must be included
in each test as well as control antigen prepared from normal (uninfected) horse RBCs.
Anticomplementary samples are examined by the IFA test. Donkey sera are frequently anticomplementary.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
No biological products are available currently.
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HIRATA H., IKADAI H., YOKOYAMA N., XUAN X., FUJISAKI K., SUZUKI N., MIKAMI T. & IGARASHI I. (2002). Cloning of a
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HOLMAN P.J., FRERICHS W.M., CHIEVES L. & WAGNER G.G. (1993). Culture confirmation of the carrier status of
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Cloning and expression of a 48-kilodalton Babesia caballi merozoite rhoptry protein and potential use of the
recombinant antigen in an enzyme-linked immunosorbent assay. J. Clin. Microbiol., 37, 3475–3480.
KAPPMEYER L.S., PERRYMAN L.E., HINES S.A., BASZLER T.V., KATZ J.B., HENNAGER S.G. & KNOWLES D.P. (1999).
Detection of equine antibodies to Babesia caballi recombinant B. caballi rhoptry-associated protein 1 in a
competitive-inhibition enzyme-linked immunosorbent assay. J. Clin. Microbiol., 37, 2285–2290.
KAPPMEYER L.S., PERRYMAN L.E. & KNOWLES D.P (1993). A Babesia equi gene encodes a surface protein with
homology to Theileria species. Mol. Biochem. Parasitol., 62, 121–124.
KATZ J., DEWALD R. & NICHOLSON J. (2000). Procedurally similar competitive immunoassay systems for the
serodiagnosis of Babesia equi, Babesia caballi, Trypanosoma equiperdum and Burkholderia mallei infection in
horses. J. Vet. Diagn. Invest., 12, 46–50.
KNOWLES D.P. KAPPMEYER, L.S. & PERRYMAN L.E. (1997). Genetic and biochemical analysis of erythrocyte-stage
surface antigens belonging to a family of highly conserved proteins of Babesia equi and Theileria species. Mol.
Biochem. Parasitol., 90, 69–79.
KNOWLES D.P., KAPPMEYER, L.S., STILLER D., HENNAGER S.G. & PERRYMAN L.E. (1992). Antibody to a recombinant
merozoite protein epitope identifies horses infected with Babesia equi. J. Clin. Microbiol., 30, 3122–3126.
KNOWLES D.P., PERRYMAN L.E. & KAPPMEYER L.S. (1991). Detection of equine antibody to Babesia equi merozoite
proteins by a monoclonal antibody-based competitive inhibition enzyme-linked immunosorbent assay. J. Clin.
Microbiol., 29, 2056–2058.
LEVINE N.D. (1985). Veterinary protozoology. Iowa State University Press, Ames, Iowa, USA.
LITTLEJOHN A. (1963). Babesiosis. In: Equine Medicine and Surgery, Bone J.F., Catcott E.J., Gabel A.A., Johnson
L.E. & Riley W.F., eds. American Veterinary Publications, California, USA, 211–220.
MADDEN P.A. & HOLBROOK A.A. (1968). Equine piroplasmosis: Indirect fluorescent antibody test for Babesia caballi.
Am. J. Vet. Res., 29, 117–123.
MAHONEY D.F. & SAAL J.R. (1961). Bovine babesiosis: Thick blood films for the detection of parasitaemia. Aust.
Vet. J., 37, 44–47.
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MCGUIRE T.C., VAN HOOSIER G.L. JR & HENSON J.B. (1971). The complement-fixation reaction in equine infectious
anemia: demonstration of inhibition by IgG (T). J. Immunol., 107, 1738–1744.
MEHLHORN H. & SCHEIN E. (1984). The piroplasms: Life cycle and sexual stages. Adv. Parasitol., 23, 37–103.
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467-475.
MORZARIA S.P., BROCKLESBY D.W. & HARRADINE D.L. (1977). Evaluation of the indirect fluorescent antibody test for
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phylogenetic analysis. Vet. Parasitol., 123, 41–54.
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routine detection of Babesia equi in horses. Vet. Parasitol., 114, 81–87.
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STILLER D. & FRERICH S.W.M. (1979). Experimental transmission of Babesia caballi to equids by different stages of
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STILLER D., FRERICHS W.M., LEATCH G. & KUTTLER K.L. (1980). Transmission of equine babesiosis and bovine
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an enzyme-linked immunosorbent assay. Clin. Diagn. Lab. Immunol., 11, 211–215.
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UNITED STATES DEPARTMENT OF AGRICULTURE (USDA) (1997). Complement fixation test for the detection of
antibodies to Babesia caballi and Babesia equi – microtitration test. USDA, Animal and Plant Health Inspection
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Services, National Veterinary Services Laboratories, Ames, Iowa, USA.
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(2001). Expression of Babesia equi merozoite antigen 1 in insect cells by recombinant baculovirus and evaluation
of its diagnostic potential in an enzyme-linked immunosorbant assay. J. Clin. Microbiol., 39, 705–709.
ZWEYGARTH E., JUST M.C. & DE WAAL D.T. (1995). Continuous in vitro cultivation of erythrocytic stages of Babesia
equi. Parasitol. Res., 81, 355-358.
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*
* *
NB: There are OIE Reference Laboratories for Equine piroplasmosis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests and reagents for equine piroplasmosis
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CHAPTER 2.5.9.
EQUINE RHINOPNEUMONITIS
SUMMARY
Equine rhinopneumonitis (ER) is a collective term for any one of several highly contagious, clinical
disease entities of equids that may occur as a result of infection by either of two closely related
herpesviruses, equid herpesvirus-1 and -4 (EHV-1 and EHV-4).
Infection by either EHV-1 or EHV-4 is characterised by a primary respiratory tract disease of
varying severity that is related to the age and immunological status of the infected animal.
Infections by EHV-1 in particular are capable of progression beyond the respiratory mucosa to
cause the more serious disease manifestations of abortion, perinatal foal death, or neurological
dysfunction.
Identification of the agent: The standard method of identification of the herpesviral agents of ER
continues to be laboratory isolation of the virus from appropriate clinical or necropsy material,
followed by seroconfirmation of its identity. The viruses can be isolated in equine cell culture from
nasopharyngeal samples taken from horses during the febrile stage of respiratory tract infection,
from liver, lung, spleen, or thymus of aborted fetuses and early foal deaths, and from the leukocyte
fraction of the blood of animals with acute EHV-1 disease. Positive identification of viral isolates as
EHV-1 or EHV-4 can be achieved by immunofluorescence with type-specific monoclonal
antibodies.
A rapid presumptive diagnosis of rhinopneumonitis abortion can be achieved by direct
immunofluorescent detection of viral antigen in cryostat sections of tissues from aborted fetuses,
using conjugated polyclonal antiserum.
Sensitive and reliable methods for EHV-1/4 detection by polymerase chain reaction or
immunoperoxidase staining have been developed and are useful adjuncts to standard virus
cultivation techniques for diagnosis of ER.
Post-mortem demonstration of histopathological lesions of EHV-1 in tissues from aborted fetuses,
cases or perinatal foal death or in the central nervous system of neurologically affected animals
complements the laboratory diagnosis of ER.
Serological tests: Because most horses will possess some level of antibody to EHV-1/4, the
demonstration of specific antibody in the serum collected from a single blood sample is not
sufficient for a positive diagnosis of recent, active ER. Paired, acute and convalescent sera from
animals suspected of being infected with EHV-1 or EHV-4 can be tested for a four-fold or greater
rise in virus-specific antibody titre by either virus neutralisation, or enzyme-linked immunosorbent
assay, or complement fixation.
Requirements for vaccines and diagnostic biologicals: Both live attenuated and inactivated
viral vaccines of varying composition are commercially available for use in assisting in the control of
ER. While vaccination is helpful in reducing the incidence of abortion in mares, and in ameliorating
the severity of clinical signs of respiratory infection in young horses, it should not be considered to
be a substitute for strict adherence to the well established tenets of sound management practices
known to reduce the risk of rhinopneumonitis. Revaccination at frequent intervals is recommended
with each of the products, as the duration of vaccine-induced immunity is relatively short.
Standards for production and licensing of both attenuated and inactivated EHV vaccines are
established by appropriate veterinary regulatory agencies in the countries of vaccine manufacture
and use. A single set of internationally recognised standards for ER vaccines is not available. In
each case, however, vaccine production is based on the system of a detailed outline of production
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employing a well characterised cell line and a master seed lot of vaccine virus that has been
validated with respect to virus identity, safety, virological purity, immunogenicity, and the absence of
extraneous microbial agents.
A. INTRODUCTION
Equine rhinopneumonitis (ER) is an historically derived term that describes a constellation of several disease
entities of horses that may include respiratory disease, abortion, neonatal foal pneumonitis, or
myeloencephalopathy (Allen & Bryans, 1986; Allen et al., 1999; Bryans & Allen, 1988; Crabb & Studdert, 1995).
The disease has been recognised for over 60 years as a threat to the international horse industry, and is caused
by either of two members of the Herpesviridae family, equid herpesvirus-1 and -4 (EHV-1 and EHV-4). EHV-1 and
EHV-4 are closely related alphaherpesviruses of horses with nucleotide sequence identity within individual
homologous genes ranging from 55% to 84%, and amino acid sequence identity from 55% to 96% (Telford et al.,
1992; 1998). The two herpesviruses are enzootic in all countries in which large populations of horses are
maintained as part of the cultural tradition or agricultural economy. There is no recorded evidence that the two
herpesviruses of ER pose any health risks to humans working with the agents.
ER is highly contagious among susceptible horses, with viral transmission to cohort animals occurring by
inhalation of aerosols of virus-laden respiratory secretions. Extensive use of vaccines has not eliminated EHV
infections, and the world-wide annual financial burden from these equine pathogens is immense.
In horses under 3 years of age, clinical ER usually takes the form of an acute, febrile respiratory illness that
spreads rapidly through the group of animals. The viruses infect and multiply in epithelial cells of the respiratory
mucosa. Signs of infection become apparent 2–8 days after exposure to virus, and are characterised by fever,
inappetence, depression, and nasal discharge. The severity of respiratory disease varies with the age of the
horse and the level of immunity resulting from previous vaccination or natural exposure. Subclinical infections with
EHV-1/4 are common, even in young animals. Although mortality from uncomplicated ER is rare and complete
recovery within 1–2 weeks is the normal pattern, the respiratory infection is a frequent and significant cause of
interrupted schedules among horses assembled for training, racing, or competitive equestrian events. Fully
protective immunity resulting from infection is of short duration, and convalescent animals are susceptible to
reinfection by EHV-1/4 after several months. Although reinfections by the two herpesviruses cause less severe or
clinically inapparent respiratory disease, the risks of subsequent abortion and/or central nervous system (CNS)
disease are not eliminated. The greatest clinical threats to individual breeding, racing, or pleasure horse
operations posed by ER are the potential abortigenic and neurological sequelae of EHV-1 respiratory infection.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent (Allen et al., 2004)
Because ER is a highly contagious disease with the potential for occurring as explosive outbreaks with high
mortality from abortigenic or neurological sequelae, rapid diagnostic methods are important. Although several
rapid and innovative diagnostic techniques based on enzyme-linked immunosorbent assay (ELISA), polymerase
chain reaction (PCR), immunohistochemical staining with peroxidase, or nucleic acid hybridisation probes have
been recently described, their use is often restricted to specialised reference laboratories, and thus the method of
choice for diagnosis of ER by diagnostic virology laboratories handling many routine samples continues to be the
traditional methodology of cell culture isolation followed by sero-identification of the isolated viruses. Successful
laboratory isolation of EHV-1/4 depends on strict adherence to proper methods for both sample collection and
laboratory processing.
a)
Collection of samples
Samples of nasopharyngeal exudate for virus isolation are best obtained from horses during the very early,
febrile stages of the respiratory disease, and are collected via the nares by swabbing the nasopharyngeal
area with a 5 × 5 cm gauze sponge attached to the end of a 50 cm length of flexible, stainless steel wire
encased in latex rubber tubing. A guarded uterine swab devise can also be used. After collection, the swab
should be removed from the wire and transported immediately to the virology laboratory in 3 ml of cold (not
frozen) fluid transport medium (serum-free MEM [minimal essential medium] with antibiotics). Virus infectivity
can be prolonged by the addition of bovine serum albumin or gelatine to 0.1% (w/v).
Virological examination of fetal tissues from suspect cases of EHV-1 abortion is most successful when
performed on aseptically collected samples of liver, lung, thymus, and spleen. The tissue samples should be
transported to the laboratory and held at 4°C until inoculated into tissue culture. Samples that cannot be
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processed within a few hours should be stored at –70°C. In ante-mortem cases of EHV-1 neurological
disease, the virus can often be isolated from the leukocyte fraction of the blood of acutely infected horses or,
less often, from the nasopharynx of the affected animal or cohort animals. For attempts at virus isolation
from blood leukocytes, a 20 ml sample of sterile blood, collected in citrate, or heparin anticoagulant (EDTA
[ethylene diamine tetra-acetic acid] should not be used as it can destroy the cell cultures). The samples
should be transported without delay to the laboratory on ice, but not frozen. Although the virus has, on
occasion, been isolated from post-mortem cases of EHV-1 neurological disease by culture of samples of
brain and spinal cord, such attempts to isolate virus are often unsuccessful; however, they maybe useful for
PCR examination.
b)
Virus isolation
For efficient primary isolation of EHV-4 from horses with respiratory disease, equine-derived cell cultures
must be used. Both EHV-1 and EHV-4 may be isolated from nasopharyngeal samples using primary equine
fetal kidney cells or cell strains of equine fibroblasts derived from dermal (E-Derm) or lung tissue. EHV-1 can
be isolated on other cell types, as will be discussed later. The nasopharyngeal swab and its accompanying
3 ml of transport medium are transferred into the barrel of a sterile 10 ml syringe. Using the syringe plunger,
the fluid is squeezed from the swab into a sterile tube. A portion of the expressed fluid is then filtered
through a sterile, 0.45 µm membrane syringe filter unit into a second sterile tube. Filtration will decrease
bacterial contamination, but may also lower virus titre. Recently prepared cell monolayers in 25 cm2 tissue
culture flasks are inoculated with 0.5 ml of the filtered, as well as the unfiltered, nasopharyngeal swab
extract. Cell monolayers in multiwell plates incubated in a 5% CO2 environment may also be used. Virus is
allowed to attach by incubating the inoculated monolayers at 37°C on a platform rocker for 1.5–2 hours.
Monolayers of uninoculated control cells should be incubated in parallel with sterile transport medium only.
At the end of the attachment period, the inocula are removed and the monolayers are rinsed twice with
phosphate buffered saline (PBS) to remove virus-neutralising antibody that may be present in the
nasopharyngeal secretions. After addition of 5 ml of supplemented maintenance medium (MEM containing
2% fetal calf serum [FCS] and twice the standard concentrations of antibiotics [penicillin, streptomycin,
gentamicin, and amphotericin B]), the flasks are incubated at 37°C. The use of positive control virus samples
to validate the isolation procedure carries the risk that this may lead to eventual contamination of diagnostic
specimens. This risk can be minimised by using routine precautions and good laboratory technique,
including the use of biosafety cabinets, inoculating positive controls after the diagnostic specimens,
decontaminating the surfaces in the hood while the inoculum is adsorbing and using a positive control of
relatively low titre. Inoculated flasks should be inspected daily by microscopy for the appearance of
characteristic herpesvirus cytopathic effect (CPE) (focal rounding, increase in refractility, and detachment of
cells). Cultures exhibiting no evidence of viral CPE after 1 week of incubation should be blind-passaged into
freshly prepared monolayers of cells, using small aliquots of both media and cells as the inoculum. Further
blind passage is usually not productive.
A number of cell types may be used for isolation of EHV-1 from the tissues of aborted fetuses or from postmortem cases of neurological disease (e.g. rabbit kidney [RK-13], baby hamster kidney [BHK-21], Madin–
Darby bovine kidney [MDBK], pig kidney [PK-15], etc.), but equine-derived cell cultures are most sensitive
and must be used if the infrequent cases of EHV-4 abortion are to be detected. Around 10% (w/v) pooled
tissue homogenates of liver, lung, thymus, and spleen (from aborted fetuses) or of CNS tissue (from cases
of neurological disease) are used for virus isolation. These are prepared by first mincing small samples of
tissue into 1 mm cubes in a sterile Petri dish with dissecting scissors, followed by macerating the tissue
cubes further in serum-free culture medium with antibiotics using a mechanical tissue grinder (e.g. TenBroeck or Stomacher). After centrifugation at 1200 g for 10 minutes, the supernatant is removed and 0.5 ml
is inoculated into duplicate cell monolayers in 25 cm2 flasks. Following incubation of the inoculated cells at
37°C for 1.5–2 hours, the inocula are removed and the monolayers are rinsed twice with PBS. After addition
of 5 ml of supplemented maintenance medium, the flasks are incubated at 37°C for up to 1 week or until
viral CPE is observed.
Culture of peripheral blood leukocytes for the presence of EHV-1 can be attempted in horses during the
early stages of the myeloencephanlopathy. Buffy coats may be prepared from unclotted blood by
centrifugation at 600 g for 15 minutes, and the buffy coat is taken after the plasma has been carefully
removed. The buffy coat is then layered onto Ficoll 1,090, centrifuged at 400 g for 20 minutes and the
leukocyte-rich interface is then layered onto Ficoll 1.077 and centrifuged in the same way. The PBMC
interface (without most granulocytes) is washed twice in PBS (300 g for 10 minutes) and resuspended in
1 ml of MEM containing 2% FCS. Then, 0.5 ml of the rinsed cell suspension is added to each of the
duplicate monolayers of equine fibroblast, equine fetal or RK-13 cell monolayers in 25 cm2 flasks containing
8–10 ml freshly added maintenance medium. The flasks are incubated at 37°C for 7 days; the inoculum is
not removed. Because CPE may be difficult to detect in the presence of the massive inoculum of leukocytes,
each flask of cells is freeze–thawed after 7 days of incubation and the contents centrifuged at 300 g for
10 minutes. Finally, 0.5 ml of the cell-free, culture medium supernatant is transferred to freshly made cell
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monolayers that are just subconfluent. These are incubated and observed for viral CPE for at least 5–6 days
before discarding as negative.
c)
Seroconfirmation of virus identity
The basis for identification of any herpesvirus isolate recovered from specimens submitted from suspected
cases of ER is its immunoreactivity with specific antisera. Specific identification of an isolate as EHV-1 or
EHV-4 can be quickly and simply accomplished by immunofluorescent (FA) detection of viral antigen in the
infected cell culture using type-specific monoclonal antibodies (MAbs), which are available from OIE
Reference Laboratories for equine rhinopneumonitis. The test, which is type-specific and accurate, can be
performed on a small aliquot of infected cells from the same container inoculated with clinical or postmortem material. An isolate made in a laboratory that lacks MAbs or FA capability can be confirmed as EHV1/4 by virus neutralization using a virus-specific polyclonal antiserum or by the PCR (see section B.1.f).
Cell monolayers infected with the isolate are removed by scraping from the flask when at least 75% CPE is
evident. The cells are pelleted from the culture medium and resuspended in 0.5 ml of PBS. 50 µl of the cell
suspension is placed into two wells of a multiwell microscope slide, air-dried, and fixed for 10 minutes with
100% acetone. Control cell suspensions (uninfected, EHV-1 infected, or EHV-4 infected) are also spotted
into each of two wells of the same slide. Control cells may be prepared in advance and stored frozen in
small aliquots. A drop of an appropriate dilution of MAb specific for EHV-1 is added to one well of each cell
pair, and a drop of MAb specific for EHV-4 is added to each of the other wells. After 30 minutes’ incubation
at 37°C in a humid chamber, unreacted antibody is removed by two 10-minute washes with PBS. MAbs
bound to viral antigen can be detected with goat anti-mouse IgG conjugated with fluorescein isothiocyanate
(FITC). A drop of diluted conjugate is added to each well and, after 30 minutes at 37°C, the wells are again
washed twice with PBS. Cells are examined with a fluorescence microscope, and positive fluorescence with
the antibody of appropriate specificity indicates the virus type.
d)
Virus detection by direct immunofluorescence
Direct immunofluorescent detection of EHV-1 antigens in samples of post-mortem tissues collected from
aborted equine fetuses provides an indispensable method to the veterinary diagnostic laboratory for making
a rapid preliminary diagnosis of herpesvirus abortion (Gunn, 1992). Side-by-side comparisons of the
immunofluorescent and cell culture isolation techniques on more than 100 cases of equine abortion have
provided evidence that the diagnostic reliability of direct immunofluorescent staining of fetal tissues obtained
at necropsy approaches that of virus isolation attempts from the same tissues. In the United States of
America (USA), specific and potent polyclonal antiserum to EHV-1, prepared in swine and conjugated with
FITC, is provided to veterinary diagnostic laboratories for this purpose by the National Veterinary Services
Laboratories of the United States Department of Agriculture (USDA). The antiserum cross-reacts with EHV-4
and hence is not useful for serotyping. Freshly dissected samples (5 × 5 mm pieces) of fetal tissue (lung,
liver, thymus, and spleen) are frozen, sectioned on a cryostat at –20°C, mounted on to microscope slides,
and fixed with 100% acetone. After air-drying, the sections are incubated at 37°C in a humid atmosphere for
30 minutes with an appropriate dilution of the conjugated swine antibody to EHV-1. Unreacted antibody is
removed by two washes in PBS, and the tissue sections are then covered with aqueous mounting media
and a cover-slip, and examined for fluorescent cells indicating the presence of EHV antigen. Each test
should include a positive and negative control consisting of sections from known EHV-1 infected and
uninfected fetal tissue.
e)
Virus detection by immunoperoxidase staining
Enzyme immunohistochemical (IH) staining methods (e.g. immunoperoxidase) have been developed
recently as procedures for detecting EHV-1 antigen in paraffin-embedded tissues of aborted equine fetuses
or neurologically affected horses (Schultheiss et al., 1993; Whitwell et al., 1992). Such ancillary IH
techniques for antigen detection may facilitate identification of the virus in archival tissue samples or in
clinical cases in which traditional laboratory methods for EHV-1 detection have been unsuccessful.
Immunoenzymatic staining for EHV-1 is particularly useful for the simultaneous evaluation of morphological
lesions and the identification of the infectious agent. Immunoperoxidase staining for EHV-1 or EHV-4 may
also be done on infected cell monolayers (van Maanen et al., 2000). Adequate controls must be included
with each immunoperoxidase test run for evaluation of both the method specificity and antibody specificity.
f)
Virus detection by polymerase chain reaction
The PCR can be used for rapid amplification and diagnostic detection of nucleic acids of EHV-1 and -4 in
clinical specimens, paraffin-embedded archival tissue, or inoculated cell cultures (Borchers & Slater, 1993;
Lawrence et al., 1994; O’Keefe et al., 1994; Varrasso et al., 2001; Wagner et al., 1992). A variety of typespecific PCR primers have been designed to distinguish between the presence of EHV-1 and EHV-4. The
correlation between PCR and virus isolation techniques for diagnosis of EHV-1 or EHV-4 is high (Varrasso
et al., 2001). Diagnosis of ER by PCR is rapid, sensitive, and does not depend on the presence of infectious
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virus in the clinical sample. It now forms an integral part of a range of diagnostic tests currently available for
ER, each with its own advantages and limitations.
For diagnosis of active infection by EHV, PCR methods are most reliable with samples from aborted fetuses
or from nasopharyngeal swabs and peripheral blood leukocytes of foals and yearlings; they are most useful
in explosive epizootics of abortion or respiratory tract disease in which a rapid identification of the virus is
critical for guiding management strategies. PCR examinations of spinal cord and brain tissue, as well as
PBMC, are important in seeking a diagnosis on a horse with neurological signs. However, the interpretation
of the amplification by PCR of genomic fragments of EHV-1 or EHV-4 in lymph nodes or trigeminal ganglia
from adult horses is complicated by the high prevalence of latent EHV-1 and EHV-4 DNA in such tissues
(Welch et al., 1992).
A simple multiplex PCR assay for simultaneous detection of both EHV-1 and EHV-4 has been described
(Wagner et al., 1992). A more sensitive protocol for nested PCR detection of EHV-1 or EHV-4 in clinical or
pathological specimens (nasal secretions, blood leukocytes, brain and spinal cord, fetal tissues, etc.) is
described here (Borchers & Slater, 1993). This procedure has been used successfully; however, the
technology in this area is changing rapidly and other simpler more sensitive techniques are becoming
available.
i)
Prepare template DNA from test specimens: Following sample homogenisation and cell (and virion)
lysis in the presence of a chaotropic salt, nucleic acids bind selectively to silica or cationic resin
substrates. Substrate-bound nucleic acids are purified in a series of rapid wash steps followed by
recovery with low-salt elution. The reagents for performing such steps for rapid nucleic acid isolation
are available in kit format from a number of commercial sources (e.g. High Pure PCR Template
Preparation Kit, Roche Molecular Biochemicals, Indianapolis, USA; QIAamp DNA Kit, Qiagen,
Valencia, USA).
ii)
Nested primer sequences specific for EHV-1 (based on those described in Borchers & Slater, 1993):
BS-1-P1 = 5’-TCT-ACC-CCT-ACG-ACT-CCT-TC-3’ (917–936)
gB1-R-2 = 5’-ACG-CTG-TCG-ATG-TCG-TAA-AAC-CTG-AGA-G-3’ (2390–2363)
BS-1-P3 = 5’-CTT-TAG-CGG-TGA-TGT-GGA-AT-3’ (1377–1396)
gB1-R-a = 5’-AAG-TAG-CGC-TTC-TGA-TTG-AGG-3’ (2147–2127)
iii)
Nested primer sequences specific for EHV-4 (Borchers & Slater, 1993):
BS-4-P1 = 5’-TCT-ATT-GAG-TTT-GCT-ATG-CT-3’ (1705–1724)
BS-4-P2 = 5’-TCC-TGG TTG-TTA-TTG-GGT-AT-3’ (2656–2637)
BS-4-P3 = 5’-TGT-TTC-CGC-CAC-TCT-TGA-CG-3’ (1857–1876)
BS-4-P4 = 5’-ACT-GCC-TCT-CCC-ACC-TTA-CC-3’ (2456–2437)
iv)
PCR conditions for first stage amplification: Specimen template DNA (1–2 ug in 2 µl) is added to a PCR
mixture (total volume of 50 µl) containing 1 × PCR buffer (50 mM KCl, 10 mM Tris/HCl, pH 9.0, 0.1%
Triton X-100), 200 µM of each deoxynucleotide triphosphate (dNTP), 2.5 mM MgCl2, 2.0 µM of each
outer primer (BS-1-P1 and gB1-R-2 for EHV-1 detection and, in a separate reaction mixture, BS-4-P1
and BS-4-P2 for EHV-4 detection) and 0.5 u Taq DNA polymerase. Cycling parameters are: initial
denaturation at 94°C for 4 minutes; 40 cycles of 94°C for 30 seconds, 60°C for 30 seconds, and 72°C
for 90 seconds; with a final extension at 72°C for 10 minutes. Separate reaction mixtures containing
either known viral DNA or no DNA (water) should be prepared and amplified as positive and negative
controls.
v)
PCR conditions for second stage (nested) amplification: Two µl of a 1/10 dilution of the first
amplification product is added to a fresh PCR mixture (total volume of 50 µl) containing 1 × PCR buffer,
200 µM of each dNTP, 2.5 mM MgCl2, 2.0 µM of each nested primer (BS-1-P3 and gB1-R-a for EHV-1
detection and, in a separate reaction mixture, BS-4-P3 and BS-4-P4 for EHV-4 detection) and 0.5 u
Taq DNA polymerase. Cycling parameters are: initial denaturation at 94°C for 4 minutes; 40 cycles of
94°C for 30 seconds, 60°C for 30 seconds, and 72°C for 1 minute; with a final extension at 72°C for
10 minutes.
vi)
Gel analysis of amplified products: 10 µl of each final amplified product, including controls, is mixed
with 2 µl of 6 × loading dye and electrophoresed on a 1.5% agarose gel in Tris/acetate or Tris-Borate
running buffer, along with a 100 base pairs (bp) DNA ladder. The gel is stained with ethidium bromide
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and viewed by UV transillumination for amplified products of either 770 bp for EHV-1 or 580 bp for
EHV-4.
g)
Histopathology
Histopathological examination of sections of formalin-fixed, paraffin-embedded tissues from aborted fetuses
or from neurologically affected horses is an essential part of the laboratory diagnosis of these two clinical
manifestations of ER. In aborted fetuses, typical herpetic intranuclear inclusion bodies present within
bronchiolar epithelium or in cells at the periphery of areas of hepatic necrosis are pathognomonic lesions for
EHV-1. The characteristic, but not pathognomonic, microscopic lesion associated with EHV-1 neuropathy is
a degenerative thrombotic vasculitis of small blood vessels in the brain or spinal cord (perivascular cuffing
and infiltration by inflammatory cells, endothelial proliferation and necrosis, and thrombus formation).
2.
Serological tests
Because of the ubiquity of the viral agents of ER and the high seroprevalence among horses in most parts of the
world, the demonstration of a negative antibody titre to EHV-1/4 by serological testing of horses designated for
export is not part of present veterinary regulations that seek to prevent international spread of infectious diseases
of horses. Serological testing can, however, be a useful adjunct procedure for assisting in the diagnosis of ER in
horses. Serodiagnosis of ER is based on the demonstration of significant increases in antibody titres in paired
sera taken during the acute and convalescent stages of the disease. The results of tests performed on sera from
a single collection date are, in most cases, impossible to interpret with any degree of confidence. The initial (acute
phase) serum sample should be taken as soon as possible after the onset of clinical signs, and the second
(convalescent phase) serum sample should be taken 2–4 weeks later. ‘Acute phase’ sera from mares after
abortion or from horses with EHV-1 neurological disease may already contain maximal titres of EHV-1 antibody,
with no increase in titres detectable in sera collected at later dates. In such cases, serological testing of paired
serum samples from clinically unaffected cohort members of the herd for rising antibody titres against EHV-1/4
may provide information useful for retrospective diagnosis of ER within the herd. Finally, the serological detection
of antibodies to EHV-1 in heart or umbilical cord blood or other fluids of equine fetuses can be of diagnostic value
in rare cases of virologically negative fetuses aborted as a result of EHV-1 infection; in some cases, the EHV 1/4
genome maybe identified from these tissues by PCR.
Serum antibody levels to EHV-1/4 may be determined by ELISA (Dutta et al., 1983), virus neutralisation (VN)
(Thomson et al., 1976), or complement fixation (CF) tests (Thomson et al., 1976). Because there are no
internationally recognised reagents or standardised techniques for performing any of the serological tests for
detection of EHV-1/4 antibody, antibody titre determinations on the same serum may differ from one laboratory to
another. Furthermore, all of the serological tests mentioned detect antibodies that are cross-reactive between
EHV-1 and EHV-4. Nonetheless, the demonstration, by any of the tests, of a four-fold or greater rise in antibody
titre to EHV-1 or EHV-4 during the course of a clinical illness provides serological confirmation of recent infection
with one of the viruses. The ELISA and CF test have the advantage that they provide results faster and do not
require cell culture facilities. Recently, a type-specific ELISA that can distinguish between antibodies to EHV-1
and EHV-4 was developed and made commercially available (Crabb et al., 1995). The microneutralisation test is
a widely used and sensitive serological assay for detecting EHV-1/4 antibody and will thus be described here.
a)
Virus neutralisation test
This serological test is most commonly performed in flat-bottom 96-well microtitre plates (tissue culture
grade) using a constant dose of virus and doubling dilutions of equine test sera. At least two replicate wells
for each serum dilution are required. Serum-free MEM is used throughout as a diluent. Virus stocks of
known titre are diluted just before use to contain 100 TCID50 (50% tissue culture infective dose) in 25 µl.
Monolayers of E-Derm or RK-13 cells are monodispersed with EDTA/trypsin and resuspended at a
concentration of 5 × 105/ml. Note that RK-13 cells can be used with EHV-1 but do not give clear CPE with
EHV-4. Antibody positive and negative control equine sera and controls for cell viability, virus infectivity, and
test serum cytotoxicity, must be included in each assay. End-point VN titres of antibody are calculated by
determining the reciprocal of the highest serum dilution that protects 100% of the cell monolayer from virus
destruction in both of the replicate wells.
A suitable test procedure is as follows:
894
i)
Inactivate test and control sera for 30 minutes in a water bath at 56°C.
ii)
Add 25 µl of serum-free MEM to all wells of the microtitre assay plates.
iii)
Pipette 25 µl of each test serum into duplicate wells of both rows A and B of the plate. The first row
serves as the serum toxicity control and the second row as the first dilution of the test. Make doubling
dilutions of each serum starting with row B and proceeding to the bottom of the plate by sequential
mixing and transfer of 25 µl to each subsequent row of wells. Six sera can be assayed in each plate.
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iv)
Add 25 µl of the appropriately diluted EHV-1 or EHV-4 virus stock to each well (100 TCID50/well)
except those of row A, which are the serum control wells for monitoring serum toxicity for the indicator
cells. Note that the final serum dilutions, after addition of virus, run from 1/4 to1/256.
v)
A separate control plate should include titration of both a negative and positive horse serum of known
titre, cell control (no virus), virus control (no serum), and a virus titration to calculate the actual amount
of virus used in the test.
vi)
Incubate the plates for 1 hour at 37°C in 5% CO2 atmosphere.
vii)
Add 50 µl of the prepared E-Derm or RK-13 cell suspension (5 × 105 cells/ml) in MEM/10% FCS to
each well.
viii) Incubate the plates for 4–5 days at 37°C in an atmosphere of 5% CO2 in air.
ix)
Examine the plates microscopically for CPE and record the results on a worksheet. Alternatively, the
cell monolayers can be scored for CPE after fixing and staining as follows: after removal of the culture
fluid, immerse the plates for 15 minutes in a solution containing 2 mg/ml crystal violet, 10% formalin,
45% methanol, and 45% water. Then, rinse the plates vigorously under a stream of running tap water.
x)
Wells containing intact cell monolayers stain blue, while monolayers destroyed by virus do not stain.
Verify that the cell control, positive serum control, and serum cytotoxicity control wells stain blue, that
the virus control and negative serum control wells are not stained, and that the actual amount of virus
added to each well is between 101.5 and 102.5 TCID50. Wells are scored as positive for neutralisation of
virus if 100% of the cell monolayer remains intact. The highest dilution of serum resulting in complete
neutralisation of virus (no CPE) in both duplicate wells is the end-point titre for that serum.
xi)
Calculate the neutralisation titre for each test serum, and compare acute and convalescent phase
serum titres from each animal for a four-fold or greater increase.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
Both live attenuated and inactivated vaccines are available as licensed, commercially prepared products for use
as prophylactic aids in reducing the burden of disease in horses caused by EHV-1/4 infection. Clinical experience
has demonstrated that none of the vaccine preparations should be relied on to provide an absolute degree of
protection from ER. Multiple doses, repeated annually, of each of the currently marketed ER vaccines are
recommended by their respective manufacturers, with vaccination schedules that vary with the particular vaccine.
Guidelines for the production of veterinary vaccines are given in Chapter 1.1.6 Principles of veterinary vaccine
production. The guidelines given here and in chapter 1.1.6 are intended to be general in nature and may be
supplemented by national and regional requirements.
At least sixteen vaccine products for ER, each containing different permutations of EHV-1, EHV-4, and the two
subtypes of equine influenza virus, are currently marketed by five veterinary biologicals manufacturers.
The clinical indications stated on the product label for use of the several available vaccines for ER are either
herpesvirus-associated respiratory disease, abortion, or both. Only four vaccine products have met the regulatory
requirements for claiming efficacy in providing protection from herpesvirus abortion as a result of successful
vaccination and challenge experiments in pregnant mares. None of the vaccine products has been conclusively
demonstrated to prevent the occurrence of neurological disease sometimes associated with EHV-1 infection.
1.
Seed management
a)
Characteristics and culture
The master seed virus (MSV) for ER vaccines must be prepared from strains of EHV-1 and/or EHV-4 that
have been positively and unequivocally identified by both serological and genetic tests. Seed virus must be
propagated in a cell line approved for equine vaccine production by the appropriate regulatory agency. A
complete record of original source, passage history, medium used for propagation, etc., shall be kept for the
master seed preparations of both the virus(es) and cell stock(s) intended for use in vaccine production.
Permanently stored stocks of both MSV and master cell stock (MCS) used for vaccine production must be
demonstrated to be pure, safe and, in the case of MSV, also immunogenic. Generally, the fifth passage from
the MSV and the twentieth passage from the MCS are the highest allowed for vaccine production. Results of
all quality control tests on master seeds must be recorded and made a part of the licensee's permanent
records.
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b)
Validation as a vaccine
i)
Purity
Tests for master seed purity include prescribed procedures that demonstrate the virus and cell seed
stocks to be free from bacteria, fungi, mycoplasmas, and extraneous viruses. Special tests must be
performed to confirm the absence of equine arteritis virus, equine infectious anaemia virus, equine
influenza virus, equine herpesvirus-2, -3, and -5, equine rhinovirus, the alphaviruses of equine
encephalomyelitis, bovine viral diarrhoea virus (BVDV – common contaminant of bovine serum), and
porcine parvovirus (PPV – potential contaminant of porcine trypsin). The purity check should also
include the exclusion of the presence of EHV-1 from EHV-4 MSV and vice versa.
ii)
Safety
Samples of each lot of MSV to be used for preparation of live attenuated ER vaccines must be tested
for safety in horses determined to be susceptible to the virulent wild-type virus, including pregnant
mares in the last 4 months of gestation. Vaccine safety must be demonstrated in a ‘safety field trial’ in
horses of various ages from three different geographical areas. The safety trial should be conducted by
independent veterinarians using a prelicensing batch of vaccine. EHV-1 vaccines making a claim for
efficacy in controlling abortion must be tested for safety in a significant number of late gestation
pregnant mares, using the vaccination schedule that will be recommended by the manufacturer for the
final vaccine product.
iii)
Immunogenicity
Tests for immunogenicity of the EHV-1/4 MSV stocks should be performed in horses on an
experimental test vaccine prepared from the highest passage level of the MSV allowed for use in
vaccine production. The test for MSV immunogenicity consists of vaccination of horses with low
antibody titres to EHV-1/4, with doses of the test vaccine that will be recommended on the final product
label. Second serum samples should be obtained and tested for significant increases in neutralising
antibody titre against the virus, 21 days after the final dose.
iv)
Efficacy
An important part of the validation process is the capacity of a prelicensing lot of the ER vaccine to
provide a significant level of clinical protection in horses from the particular disease manifestation of
EHV-1/4 infection for which the vaccine is offered, when used under the conditions recommended by
the manufacturer's product label. Serological data are not acceptable for establishing the efficacy of
vaccines for ER. Efficacy studies must be designed to ensure appropriate randomisation of test
animals to treatment groups, blinding of the recording of clinical observations, and the use of sufficient
numbers of animals to permit statistical evaluation for effectiveness in prevention or reduction of the
specified clinical disease. The studies should be performed on fully formulated experimental vaccine
products (a) produced in accordance with, (b) at or below the minimum antigenic potency specified in,
and, (c) produced with the highest passage of MSV and MCS allowed by the approved ‘Outline of
Production’ (see Section C.2). Vaccine efficacy is demonstrated by vaccinating a minimum of 20 EHV1/4-susceptible horses possessing serum neutralising antibody titres ≤32, followed by challenge of the
vaccinates and ten nonvaccinated control horses with virulent virus. A significant difference in the
clinical signs of ER must be demonstrated between vaccinates and nonvaccinated control horses. The
vaccination and challenge study must be performed on an identical number of pregnant mares and
scored for abortion if the vaccine product will make a label usage claim ‘for prevention of’ or ‘as an aid
in the prevention of’ abortion caused by EHV-1.
2.
Method of manufacture
A detailed protocol of the methods of manufacture to be followed in the preparation of vaccines for ER must be
compiled, approved, and filed as an Outline of Production with the appropriate licensing agency. Specifics of the
methods of manufacture for ER vaccines will differ with the type (live or inactivated) and composition (EHV-1 only,
EHV-1 and EHV-4, EHV-4 and equine influenza viruses, etc.) of each individual product, and also with the
manufacturer.
3.
In-process control
Cells, virus, culture medium, and medium supplements of animal origin that are used for the preparation of
production lots of vaccine must be derived from bulk stocks that have passed the prescribed tests for bacterial,
fungal, and mycoplasma sterility; nontumorgenicity; and absence of extraneous viral agents.
4.
Batch control
Each bulk production lot of ER vaccine must pass tests for sterility, safety, and immunogenic potency.
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a)
Sterility
Samples taken from each batch of completed vaccine are tested for bacteria, fungi, and mycoplasma
contamination. Procedures to establish that the vaccine is free from extraneous viruses are also required;
such tests should include inoculation of cell cultures that allow detection of the common equine viruses, as
well as techniques for the detection of BVDV and PPV in ingredients of animal origin used in the production
of the batch of vaccine.
b)
Safety
Tests to assure safety of each production batch of ER vaccine must demonstrate complete inactivation of
virus (for inactivated vaccines) as well as a level of residual virus-killing agent that does not exceed the
maximal allowable limit (e.g. 0.2% for formaldehyde). Safety testing in laboratory animals is also required.
c)
Potency
Batch control of antigenic potency for EHV-1 vaccines only may be tested by measuring the ability of
dilutions of the vaccine to protect hamsters from challenge with a lethal dose of hamster-adapted EHV-1
virus. Although potency testing on production batches of ER vaccine may also be performed by vaccination
of susceptible horses followed by either viral challenge or assay for seroconversion, the recent availability of
virus type-specific MAbs has permitted development of less costly and more rapid in-vitro immunoassays for
antigenic potency. The basis for such in-vitro assays for ER vaccine potency is the determination, by use of
the specific MAb, of the presence of at least the minimal amount of viral antigen within each batch of vaccine
that correlates with the required level of protection (or seroconversion rate) in a standard animal test for
potency.
d)
Duration of immunity
Tests to establish the duration of immunity to EHV-1/4 achieved by immunisation with each batch of vaccine
are not required. The results of many reported observations indicate that vaccination-induced immunity to
EHV-1/4 is not more than a few months in duration; these observations are reflected in the frequency of
revaccination recommended on ER vaccine product labels.
e)
Stability
At least three production batches of vaccine should be tested for shelf life before reaching a conclusion on
the vaccine’s stability. When stored at 4°C, inactivated vaccine products generally maintain their original
antigenic potency for at least 1 year. Lyophilised preparations of the live virus vaccine are also stable during
storage for 1 year at 4°C. Following reconstitution, live virus vaccine is unstable and cannot be stored
without loss of potency.
5.
Tests on the final product
Before release for labelling, packaging, and commercial distribution, randomly selected filled vials of the final
vaccine product must be tested by prescribed methods for freedom from contamination and safety in laboratory
test animals.
a)
Safety
See Section C.4.b.
b)
Potency
See Section C.4.c.
REFERENCES
ALLEN G.P. & BRYANS J.T. (1986). Molecular epidemiology, pathogenesis and prophylaxis of equine herpesvirus-1
infections. In: Progress in Veterinary Microbiology and Immunology, Vol. 2, Pandey R., ed. Karger, Basel,
Switzerland & New York, USA, 78–144.
ALLEN G.P., KYDD J.H., SLATER J.D. & SMITH K.C. (1999). Recent advances in understanding the pathogenesis,
epidemiology, and immunological control of equid herpesvirus-1 (EHV-1) abortion. Equine Infect. Dis., 8, 129–
146.
ALLEN G.P., KYDD J.H., SLATER J.D. & SMITH K.C. (2004). Equid herpesvirus-1 (EHV-1) and -4 (EHV-4) infections.
In: Infectious Diseases of Livestock, Coetzer J.A.W., Thomson G.R. & Tustin R.C., eds. Oxford University Press,
Cape Town, South Africa.
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BORCHERS K. & SLATER J. (1993). A nested PCR for the detection and differentiation of EHV-1 and EHV-4. J. Virol.
Methods, 45, 331–336.
BRYANS J.T. & ALLEN G.P. (1988). Herpesviral diseases of the horse. In: Herpesvirus Diseases of Animals,
Wittman G., ed. Kluwer, Boston, USA, 176–229.
CRABB B.S., MACPHERSON C.M., REUBEL G.H., BROWNING G.F., STUDDERT M.J. & DRUMMER H.E. (1995). A typespecific serological test to distinguish antibodies to equine herpesviruses 4 and 1. Arch. Virol., 140, 245–258.
CRABB B.S. & STUDDERT M.J. (1995). Equine herpesviruses 4 (equine rhinopneumonitis virus) and 1 (equine
abortion virus). Adv. Virus Res., 45, 153–190.
DUTTA S.K., TALBOT N.C. & MYRUP A.C. (1983). Detection of equine herpesvirus-1 antigen and the specific
antibody by enzyme-linked immunosorbent assay. Am. J. Vet. Res., 44, 1930–1934.
GUNN H.M. (1992). A direct fluorescent antibody technique to diagnose abortion caused by equine herpesvirus.
Irish Vet. J., 44, 37–40.
LAWRENCE G.L., GILKERSON J., LOVE D.N., SABINE M. & WHALLEY J.M. (1994). Rapid, single-step differentiation of
equid herpesvirus 1 and 4 from clinical material using the polymerase chain reaction and virus-specific primers. J.
Virol. Methods, 47, 59–72.
O’KEEFE J.S., JULIAN A., MORIARTY K., MURRAY A. & WILKS C.R. (1994). A comparison of the polymerase chain
reaction with standard laboratory methods for the detection of EHV-1 and EHV-4 in archival tissue samples. N.Z.
Vet. J., 42, 93–96.
SCHULTHEISS P.C., COLLINS J.K. & CARMAN J. (1993). Use of an immunoperoxidase technique to detect equine
herpesvirus-1 antigen in formalin-fixed paraffin-embedded equine fetal tissues. J. Vet. Diagn. Invest., 5, 12–15.
TELFORD E.A.R., WATSON M.S., MCBRIDE K. & DAVISON A.J. (1992). The DNA sequence of equine herpesvirus-1.
Virology, 189, 304–316.
TELFORD E.A.R., WATSON M.S., PERRY J., CULLINANE A.A. & DAVISON A.J. (1998). The DNA sequence of equine
herpesvirus 4. J. Gen. Virol., 79, 1197–1203.
THOMSON G.R., MUMFORD J.A., CAMPBELL J., GRIFFITHS L. & CLAPHAM P. (1976). Serological detection of equid
herpesvirus 1 infections of the respiratory tract. Equine Vet. J., 8, 58–65.
MAANEN C., VREESWIJK J., MOONEN P., BRINKHOF J. DE BOER-LUIJTZE E. & TERPSTRA C. (2000). Differentiation
and genomic and antigenic variation among fetal, respiratory, and neurological isolates from EHV1 and EHV4
infections in The Netherlands. Vet. Q., 22 (2): 88–93.
VAN
VARRASSO A., DYNON K., FICORILLI N., HARTLEY C.A., STUDDERT M.J. & DRUMMER H.E. (2001). Identification of
equine herpesviruses 1 and 4 by polymerase chain reaction. Aust. Vet. J., 79, 563–569.
WAGNER W.N., BOGDAN J., HAINES D., TOWNSEND H.G.G. & MISRA V. (1992). Detection of equine herpesvirus and
differentiation of equine herpesvirus type 1 from type 4 by the polymerase chain reaction. Can. J. Microbiol., 38,
1193–1196.
WELCH H.M., BRIDGES C.G., LYON A.M., GRIFFITHS L. & EDINGTON N. (1992). Latent equid herpesviruses 1 and 4:
detection and distinction using the polymerase chain reaction and cocultivation from lymphoid tissues. J. Gen.
Virol., 73, 261–268.
WHITWELL K.E., GOWER S.M. & SMITH K.C. (1992). An immunoperosidase method applied to the diagnosis of
equine herpesvirus abortion, using conventional and rapid microwave techniques. Equine Vet. J., 24, 10–12.
*
* *
NB: There are OIE Reference Laboratories for Equine rhinopneumonitis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for equine rhinopneumonitis
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.10.
EQUINE VIRAL ARTERITIS
SUMMARY
Equine viral arteritis (EVA) is a contagious viral disease of equids caused by equine arteritis virus
(EAV), an RNA virus classified in the family Arteriviridae. Only one major serotype of the virus has
been identified so far. Equine arteritis virus is found in horse populations in many countries worldwide. Although infrequently reported in the past, confirmed outbreaks of EVA appear to be on the
increase.
The majority of naturally acquired infections with EAV are subclinical. Where present, clinical signs
of EVA can vary in range and severity. The disease is characterised principally by fever,
depression, anorexia, dependent oedema, especially of the limbs, scrotum and prepuce in the
stallion, conjunctivitis, an urticarial-type skin reaction, abortion and, rarely, a fulminating pneumonia,
enteritis or pneumo-enteritis in young foals. Apart from mortality in young foals, the case-fatality
rate in outbreaks of EVA is very low. Affected horses almost invariably make complete clinical
recoveries. A long-term carrier state can occur in a variable percentage of infected stallions, but not
in mares, geldings or sexually immature colts.
Identification of the agent: EVA cannot be differentiated clinically from a number of other
respiratory and systemic equine diseases. Diagnosis of EAV infection is based on virus isolation,
detection of nucleic acid or viral antigen, or demonstration of a specific antibody response. Virus
isolation should be attempted from appropriate clinical or post-mortem specimens in rabbit, equine,
or monkey kidney cell culture. The identity of isolates of EAV should be confirmed by neutralisation
test, reverse-transcription polymerase chain reaction (RT-PCR) assay, or by immunocytochemical
methods, namely indirect immunofluorescence or avidin–biotin–peroxidase techniques.
Detection and identification of EAV nucleic acid in suspect cases of the disease can also be
attempted using the RT-PCR assay and appropriate viral-specific RNA primers.
Where mortality is associated with a suspected outbreak of EVA, a wide range of tissues should be
examined for histological evidence of panvasculitis that is especially pronounced in the small
arteries throughout the body. The characteristic vascular lesions present in the mature animal are
not a notable feature in EVA-related abortions. In such cases, equine arteritis viral antigens may be
visualised by immunohistochemical examination of placental and various fetal tissues.
Serological tests: A variety of serological tests, including virus neutralisation (VN), complement
fixation (CF), indirect fluorescent antibody, agar gel immunodiffusion, the enzyme-linked
immunosorbent assay (ELISA), and the fluorescent microsphere immunoassay assay (MIA) have
been used for the detection of antibody to EAV. The tests currently in widest use are the
complement-enhanced VN test and the ELISA. The VN test is a very sensitive and highly specific
assay of proven value in diagnosing acute infection and in seroprevalence studies. Several ELISAs
have been developed, none of which have been as extensively validated as the VN test though
some appear to offer comparable specificity and close to equivalent sensitivity. The CF test is less
sensitive than either procedure, but it can be used for diagnosing recent infection.
Requirements for vaccines and diagnostic biologicals: Two commercial tissue culture vaccines
are currently available against EVA. One is a modified live virus (MLV) vaccine prepared from virus
that has been attenuated for horses by multiple serial transfers in primary equine and rabbit cell
cultures. It has been shown to be safe and protective for stallions and nonpregnant mares.
Vaccination of foals under 6 weeks of age and of pregnant mares in the final 2 months of gestation
is contraindicated. There is no evidence of back reversion to virulence of the vaccine virus following
its use in the field over more than 20 years. The second vaccine is an inactivated, adjuvanted
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product prepared from virus grown in equine cell culture that can be used in nonbreeding and
breeding horses. In the absence of appropriate safety data, the vaccine is not currently
recommended for use in pregnant mares.
A. INTRODUCTION
Equine viral arteritis (EVA) is a contagious viral disease of equids caused by equine arteritis virus (EAV), a
positive-sense, single-stranded RNA virus, and the prototype member of the genus Arterivirus, family
Arteriviridae, order Nidovirales (Cavanagh, 1997). Epizootic lymphangitis pinkeye, fièvre typhoide and
rotlaufseuche are some of the descriptive terms used in the past to refer to a disease that clinically very closely
resembled EVA. The natural host range of EAV would appear to be restricted to equids, although very limited
evidence would suggest it may also include new world camelids, viz. alpacas and llamas (Weber et al., 2006).
The virus does not present a human health hazard (Timoney & McCollum, 1993). EAV is present in the horse
population of many countries world-wide (Timoney & McCollum, 1993). There has been an increase in the
incidence of EVA in recent years that has been linked to the greater frequency of movement of horses and use of
transported semen (Balasuriya et al., 1998).
While the majority of cases of acute infection with EAV are subclinical, certain strains of the virus can cause
disease of varying severity (Timoney & McCollum, 1993). Typical cases of EVA can present with all or any
combination of the following clinical signs: fever, depression, anorexia, leukopenia, dependent oedema, especially
of the limbs, scrotum and prepuce of the stallion, conjunctivitis, ocular discharge, supra or periorbital oedema,
rhinitis, nasal discharge, a local or generalised urticarial skin reaction, abortion, stillbirths and, rarely, a fulminating
pneumonia, enteritis or pneumo-enteritis in young foals. Regardless of the severity of clinical signs, affected
horses almost invariably make complete recoveries. The case-fatality rate in outbreaks of EVA is very low;
mortality is usually only seen in very young foals, particularly those congenitally infected with the virus (Timoney &
McCollum, 1993; Vaala et al., 1992), and very rarely in otherwise healthy adult horses.
EVA cannot be differentiated clinically from a number of other respiratory and systemic equine diseases, the most
common of which are equine influenza, equine herpesvirus 1 and 4 infections, infection with equine rhinitis A and
B viruses, equine adenoviruses and streptococcal infections, with particular reference to purpura haemorrhagica.
The disease also has clinical similarities to equine infectious anaemia, African horse sickness fever, cases of
Hendra virus infection, Getah virus infection and toxicosis caused by hoary alyssum (Berteroa incana) (Timoney &
McCollum, 1993). After infection, EAV replicates in macrophages and circulating monocytes (Del Piero, 2000) and
is shed in various secretions/excretions of acutely infected animals, in especially high concentration from the
respiratory tract (McCollum et al., 1971).
A variable percentage of acutely infected stallions later become long-term carriers in the reproductive tract and
constant semen shedders of the virus (Timoney & McCollum, 1993; 2000). The carrier state, which has been
shown to be androgen dependant, has been found in the stallion, but not in the mare, gelding or sexually
immature colt (Timoney & McCollum, 1993). Unequivocal evidence of the carrier state has only been found in
stallions serologically positive for antibodies to the virus (Timoney & McCollum, 2000). While temporary downregulation of circulating testosterone levels using a GnRH antagonist or by immunisation with GnRH would
appear to have expedited clearance of the carrier state in some stallions, the efficacy of either treatment strategy
has yet to be fully established. Concern has been expressed that such a therapeutic approach could be used to
deliberately mask existence of the carrier state.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
a)
Virus isolation
Where an outbreak of EVA is suspected, or when attempting to confirm a case of subclinical EAV infection,
virus isolation should be attempted from nasopharyngeal and conjunctival swabs, unclotted blood samples,
and semen from stallions considered to be possible carriers of the virus (Timoney & McCollum, 1993). To
optimise the chances of virus isolation, the relevant specimens should be obtained as soon as possible after
the onset of fever in affected horses. As heparin can inhibit the growth of EAV in rabbit kidney cells (RK-13
cell line), its use as an anticoagulant is contraindicated as it may interfere with isolation of the virus from
whole blood. Acid citrate dextrose or ethylenediaminetetraacetic acid (EDTA) are the anticoagulants of
choice to use in obtaining unclotted blood samples. Where EVA is suspected in cases of mortality in young
foals or older animals, isolation of EAV can be attempted from a variety of tissues, especially the lymphatic
glands associated with the alimentary tract and related organs, and also the lungs, liver and spleen
(McCollum et al., 1971). In outbreaks of EVA-related abortion and/or cases of stillborn foals, placental and
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fetal fluids and a wide range of placental, lymphoreticular and other fetal tissues (especially lung) can be
productive sources of virus (Timoney & McCollum, 1993).
Swabs for attempted isolation should be immersed in a suitable viral transport medium and these, together
with any fluids or tissues collected for virus isolation and/or reverse-transcription polymerase chain reaction
(RT-PCR) testing should be shipped either refrigerated or frozen in an insulated container to the laboratory,
preferably using an overnight delivery service. Unclotted blood samples must be transported refrigerated but
not frozen. Where possible, specimens should be submitted to a laboratory with established competency to
test for this infection.
Although reportedly not always successful in natural cases of EAV infection (Timoney & McCollum, 1993),
virus isolation should be attempted from clinical specimens or necropsied tissues using rabbit, equine or
monkey kidney cell culture (Timoney et al., 2004; Timoney & McCollum, 1993). Selected cell lines, e.g. RK13 (ATCC CCL-37), LLC-MK2 (ATCC CCL-7), and primary horse or rabbit kidney cell culture can be used,
with RK-13 cells being the cell system of choice (Timoney et al., 2004). Experience over the years has
shown that primary isolation of EAV from semen can present more difficulty than from other clinical
specimens or from infected tissues unless an appropriate cell culture system is used. Several factors have
been shown to influence primary isolation of EAV from semen in RK-13 cells. Higher isolation rates have
been obtained using 3- to 5-day-old monolayers, a large inoculum size in relation to the cell surface area in
the inoculated flasks or multiwell plates, and most importantly, the incorporation of carboxymethyl cellulose
(medium viscosity, 400–800 cps) in the overlay medium. It should be noted that most RK-13 cells, including
ATCC CCL-37, are contaminated with bovine viral diarrhoea virus, the presence of which appears to
enhance sensitivity of this cell system for the primary isolation of EAV, especially from semen. There is
considerable evidence to indicate that primary isolation rates of EAV may be increased by using RK-13 cells
of high passage history1 (Timoney et al., 2004).
Inoculated cultures are examined daily for the appearance of viral cytopathic effect (CPE), which is usually
evident within 2–6 days. In the absence of visible CPE, culture supernatants should be subinoculated on to
confluent cell monolayers after 4–7 days. While the vast majority of isolations of EAV are made on the first
passage in cell culture, a small minority only become evident on the second or subsequent passages in vitro
(Timoney & McCollum, 1993; 2000). The identity of isolates of EAV can be confirmed in a one-way
neutralisation test, by standard RT-PCR or real-time RT-PCR assay (Balasuriya et al., 1998) or by an
immunocytochemical method (Little et al., 1995), indirect immunofluorescence (Crawford & Henson, 1973)
or the avidin–biotin–peroxidase (ABC) technique (Little et al., 1995). A polyclonal rabbit antiserum has been
used to identify EAV in infected cell cultures. Mouse monoclonal antibodies (MAbs) to the nucleocapsid
protein (N) (Balasuriya et al., 1998) and major envelope glycoprotein (GP5) of EAV (Balasuriya et al., 1998)
and a monospecific rabbit antiserum to the unglycosylated envelope protein (M) (Balasuriya et al., 1998)
have also been developed and these can detect various strains of the virus in RK-13 cells as early as 12–
24 hours after infection (Balasuriya et al., 1998; Little et al., 1995).

Virus isolation from semen (a prescribed test for international trade)
There is considerable evidence that short- and long-term carrier stallions shed EAV constantly in the semen,
but not in respiratory secretions or urine; nor has it been demonstrated in the buffy coat (peripheral blood
mononuclear cells) of such animals (Timoney & McCollum, 1993; 2000). Stallions should first be blood
tested using the virus neutralisation (VN) test or an appropriately validated enzyme-linked immunosorbent
assay (ELISA) or other serological test procedure. Virus isolation should be attempted from the semen of
stallions serologically positive (titre ≥1/4) for antibodies to EAV that do not have a certified history of
vaccination against EVA with confirmation that they were serologically negative (titre <1/4) at time of initial
vaccination. Virus isolation is also indicated in the case of shipped semen where the serological status and
possible vaccination history of the donor stallion is not available. It is recommended that virus isolation from
semen be attempted from two samples, which can be collected on the same day, on consecutive days or
after an interval of several days or weeks. There is no evidence that the outcome of attempted virus isolation
from particular stallions is influenced by the frequency of sampling, the interval between collections or time
of the year. Isolation of EAV should be carried out preferably on portion of an entire ejaculate collected using
an artificial vagina or a condom and a teaser or phantom mare. When it is not possible to obtain semen by
this means, a less preferable alternative is to collect a dismount sample at the time of breeding. Care should
be taken to ensure that no antiseptics/disinfectants are used in the cleansing of the external genitalia of the
stallion prior to collection. Samples should contain the sperm-rich fraction of the ejaculate with which EAV is
associated as the virus is not present in the pre-sperm fraction of semen (Timoney & McCollum, 1993;
2000). Immediately following collection, the semen should be refrigerated on crushed ice or on freezer packs
for transport to the laboratory with a minimum of delay. Where there is likely to be a delay in submitting a
specimen for testing, the semen can be frozen at or below –20°C for a short period before being dispatched
1
Such a line (RK-13-KY) is available from the OIE Reference Laboratory for Equine viral arteritis at the Maxwell H. Gluck
Equine Research Center, University of Kentucky, United States of America.
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to the laboratory. Freezing a sample has not been found to militate against isolation of EAV from the semen
of a carrier stallion. In situations where it is not feasible to determine the carrier status of a stallion by virus
isolation or RT-PCR procedures, the stallion can be test bred to two seronegative mares, which are checked
for seroconversion to the virus 28 days after breeding (Timoney & McCollum, 1993).

Test procedure
i)
On receipt in the laboratory, it should be noted whether each semen sample is frozen, chilled or at
ambient temperature. Every sample should be checked to ensure that it contains the sperm-rich
fraction of the ejaculate. This can be established by microscopic examination of a wet-mount
preparation of a sample. Additionally, specimens of ejaculate should be visually inspected for colour
and presence of gross particulate contamination. If a semen specimen is contaminated with blood,
which can result from trauma to the external genitalia of the stallion at time of collection, a repeat
sample should be requested as testing such a specimen from a serologically positive stallion may
compromise the reliability of the virus isolation result.
ii)
Although no longer considered an essential step, pretreatment of semen before inoculation into cell
culture by short-term sonication (for three 15-second cycles); facilitates effective mixing and dispersion
of a sample.
iii)
After removal of culture medium, 3- to 5-day-old confluent monolayer cultures of RK-13 cells, either in
25 cm2 tissue culture flasks or multiwell plates, are inoculated with serial decimal dilutions (10–1 to
10–3) of seminal plasma in tissue culture maintenance medium containing 2% fetal bovine serum and
antibiotics. An inoculum of 1 ml per 25 cm2 flask is used and no fewer than two flasks per dilution of
seminal plasma are inoculated. Inoculum size and number of wells inoculated per dilution of a
specimen should be pro-rated where multiwell plates are used. Appropriate dilutions of a virus positive
control semen sample or virus control of known titre diluted in culture medium should be included in
each test.
iv)
The flasks are closed, lids replaced on multiwell plates and inoculated cultures gently rotated to
disperse the inoculum over the cell monolayers.
v)
Inoculated cultures are then incubated for 1 hour at 37°C either in an aerobic incubator or an incubator
containing a humidified atmosphere of 5% CO2 in air, depending on whether flasks or multiwell plates
are used.
vi)
Without removing any of the inoculum or washing the cell monolayers, the latter are overlaid with
0.75% carboxymethyl cellulose containing medium with antibiotics.
vii)
The flasks or plates are reincubated at 37°C and checked microscopically for viral CPE, which is
usually evident within 2–6 days.
viii) In the absence of visible CPE, culture supernatants are subinoculated onto 3- to 5-day-old confluent
cell monolayer cultures of RK-13 cells after 5–7 days. After removal of the overlay medium,
monolayers are stained with 0.1% formalin-buffered crystal violet solution.
The identity of any virus isolates should be confirmed by VN, immunofluorescence (Crawford & Henson,
1973) or ABC technique, using a monospecific antiserum to EAV or MAbs to the structural proteins, N or
GP5 of the virus (Balasuriya et al., 1998; Del Piero, 2000; Little et al., 1995), or by standard RT-PCR
(Balasuriya et al., 1998; Gilbert et al., 1997) or real-time RT-PCR assay (Balasuriya et al., 2002; Lu et al.,
2007; Westcott et al., 2003).
In the one-way neutralisation test, serial decimal dilutions of the virus isolate are tested against a
neutralising MAb or monospecific antiserum prepared against the prototype Bucyrus strain of EAV (ATCC
VR 796) and also a serum negative for neutralising antibodies to the virus. Corresponding titrations of the
prototype Bucyrus virus with the same reference antibody reagents are included as test controls. The test is
performed in either 25 cm2 tissue culture flasks or multiwell plates. Appropriate quantities of the known EAV
positive and negative antibody reagents are inactivated for 30 minutes in a water bath at 56°C and diluted
1/4 in phosphate buffered saline, pH 7.2; then 0.3 ml of diluted antibody reagent is dispensed into five tubes
for each virus isolate to be tested. Serial decimal dilutions (10–1 to 10–5) of each virus are made in Eagles
Minimal Essential Medium containing 10% fetal bovine serum, antibiotics and 10% freshly diluted guinea-pig
complement. Then, 0.3 ml of each virus dilution is added to the tubes containing positive and negative
antibody reagents. The tubes are shaken and the virus/antibody mixtures are incubated for 1 hour at 37°C.
The mixtures are then inoculated onto 3- to 5-day-old confluent monolayer cultures of RK-13 cells, either in
25 cm2 flasks or multiwell plates, using two flasks or wells per virus dilution. Each flask is inoculated with
0.25 ml of virus/antibody mixture; the inoculum size is pro-rated where multiwell plates are used. Inoculated
flasks or plates are incubated for 2 hours at 37°C, gently rocking after 1 hour to disperse the inoculum over
the cell monolayers. Without removing any of the inoculum or washing the cell monolayers, the latter are
overlaid with 0.75% carboxymethyl cellulose containing medium and incubated for 4–5 days at 37°C, either
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in an aerobic incubator or an incubator containing a humidified atmosphere of 5% CO2 in air. After removal
of the medium, monolayers are stained with 0.1% formalin-buffered crystal violet solution. Plaques are
counted and the virus infectivity titre is determined both in the presence and absence of EAV antibodies
using the Spearman–Kärber method. Confirmation of the identity of an isolate is based on a reduction in
plaque count of at least 102 logs of virus in the presence of antibody positive serum against the Bucyrus
strain of EAV.
The vast majority of EAV isolates from carrier stallions are made in the first passage in cell culture using the
described test procedure (Timoney & McCollum, 1993; 2000). The occurrence of nonviral cytotoxicity or
bacterial contamination of specimens is not considered significant problems when attempting isolation of this
virus from stallion semen. Nonviral cytotoxicity, if observed, usually affects monolayers inoculated with the
10–1 and, much less frequently, the 10–2 dilution of seminal plasma. Treatment of seminal plasma with
polyethylene glycol (Mol. wt 6000) prior to inoculation has been used with some success in overcoming this
problem (Fukunaga et al., 2000). The method described involves the addition of polyethylene glycol to the
10–1 to 10–3 dilutions of seminal plasma to give a final concentration of 10% in each dilution. The mixtures
are held overnight at 4°C with gentle stirring, after which they are centrifuged at 2000 g for 30 minutes and
the supernatants are discarded. The precipitates are suspended in cell culture maintenance medium to onetenth the volume of the original dilutions and the mixtures are homogenised. They are then centrifuged at
2000 g for 30 minutes and the supernatants are taken off and used for inoculation. There is no evidence to
indicate that pretreatment of seminal plasma in this manner reduces sensitivity of the virus isolation
procedure (Fukunaga et al., 2000). Where bacterial contamination of a sample is a problem, it is preferable
to request a repeat semen collection from the individual stallion. If this is not possible, an attempt can be
made to control the contamination by pre-treatment of the sample with antibiotic containing viral transport
medium, holding overnight at 4°C followed by ultracentrifugation and resuspension of the pellet before
diluting and inoculating the specimen into cell culture.
The presence of anti-EAV antibody activity in the seminal plasma of certain virus-shedding stallions has not
been found to prevent detection of the carrier state in these animals.
b)
Nucleic acid recognition methods
The standard two-step RT-PCR, single-step RT-PCR and real-time RT-PCR (rRT-PCR) assays are gaining
greater acceptance and being more widely used as an alternative to virus isolation in cell culture for the
detection of EAV in diagnostic materials. The RT-PCR-based assays provide a means of identifying virusspecific RNA in clinical specimens, namely nasopharyngeal swab filtrates, buffy coats, raw and extended
semen and urine, and in post-mortem tissue samples (Balasuriya et al., 2002; Gilbert et al., 1997; Westcott
et al., 2003). Standard two-step RT-PCR, single-step RT-PCR, RT-nested PCR (RT-nPCR), and one tube
TaqMan® rRT-PCR assays have been developed and evaluated for the detection of various strains of the
virus in tissue culture fluid, semen and nasal secretions (Balasuriya et al., 2002; Gilbert et al., 1997; Lu et
al., 2007; Westcott et al., 2003). These assays targeted six different open reading frames (ORFs) in the EAV
genome (ORFs 1b, 3–7). However, there is considerable variation in the sensitivity and specificity among
RT-PCR assays incorporating different primer pairs targeting various ORFs. Results comparable to virus
isolation have been obtained with some but not all standard single-step RT-PCR, two-step RT-PCR, RTnPCR or one tube TaqMan® rRT-PCR assays (Balasuriya et al., 2002; Gilbert et al., 1997; Lu et al., 2007).
Compared with traditional virus isolation, these RT-PCR-based assays are frequently more sensitive, less
expensive and considerably more rapid to perform, the majority taking less than 24 hours to complete. In
addition, RT-PCR assays have the advantage of not requiring viable virus for performance of the test. The
one-tube rRT-PCR assay for EAV provides a simple, rapid and reliable method for the detection and
identification of viral nucleic acid in equine semen and tissue culture fluid (Balasuriya et al., 2002; Lu et al.,
2007). The one tube rRT-PCR has the following important advantages over the standard two-step RT-PCR:
1) eliminating the possibility of cross contamination between samples with previously amplified products as
the sample tube is never opened; and 2) reducing the chance of false-positive reactions because the rRTPCR product is detected with a sequence-specific probe. Because of the high sensitivity of the RT-PCR
assay, however, and in the absence of appropriate safeguards in the laboratory, there is the potential for
cross-contamination between samples, giving rise to false-positive results. For example, the RT-nPCR
assay, while it provides enhanced sensitivity for the detection of EAV, it also increases the likelihood of
false-positive results. The risk of cross-contamination is greater using the RT-nPCR assay because of the
second PCR amplification step involving the product from the first RT-PCR reaction. To minimise the risk of
cross-contamination, considerable care needs to be taken, especially during the steps of RNA extraction and
reaction setup. Relevant EAV positive and negative template controls and, where appropriate, nucleic acid
extracted from the tissue culture fluid of uninfected cells, need to be included in each RT-PCR assay. Thus,
in many circumstances, use of the single-step RT-PCR or the one tube rRT-PCR assay would largely
circumvent the problems associated with cross contamination.
Primer selection is critical to the sensitivity of the RT-PCR assay with primers (and probe in the case of the
rRT-PCR assay) preferably designed from the most conserved region(s) of the EAV genome. Comparative
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nucleotide sequence analysis has shown that ORF 1b (encodes the viral polymerase), ORF 6 (M protein)
and 7 (N protein) are more conserved than the other ORFs among EAV strains so far analysed from North
America and Europe (Balasuriya et al., 2004; Lu et al., 2007; Westcott et al., 2003). The most conserved
gene among different strains of EAV is ORF7 and primers specific for ORF7 (and probe for rRT-PCR) have
detected a diversity of strains of the virus of European and North American origin (Balasuriya et al., 2002; Lu
et al., 2007). Furthermore, the use of multiple primer pairs specific for different ORFs 1b ([forward: 5’-GATGTC-TAT-GCT-CCA-TCA-TT-3’ and reverse: 5’-GGC-GTA-GGC-TCC-AAT-TGA-A-3’]) and/or [forward: 5’CCT-GAG-ACA-CTG-AGT-CGC-GT-3’ and reverse 5’-CCT-GAT-GCC-ACA-TGG-AAT-GA-3’]) (Gilbert et
al., 1997), ORF 6 ([forward: 5’-CTG-AGG-TAT-GGG-AGC-CAT-AG-3’ and reverse: 5’-GCA-GCC-AAA-AGCACA-AAA-GC-3’]) and ORF 7 ([forward 5’-ATG-GCG-TCA-AGA-CGA-TCA-CG-3’ and reverse 5’-AGA-ATATCC-ACG-TCT-TAC-GGC-3’]) markedly increases the likelihood of detecting North American and European
strains of EAV in the RT-PCR assay. The two primer pairs specific for ORF 1b are suitable for use in the RTrPCR assay (Gilbert et al., 1997). While the RT-PCR has been found to be highly sensitive for viral nucleic
acid detection in raw semen, there is evidence to that it is not of equivalent reliability when testing extended
or cryopreserved semen of very low virus infectivity.
In addition to the foregoing RT-PCR assays, 2 TaqMan® fluorogenic probe-based one-tube rRT-PCR
assays have been described for the detection of EAV nucleic acid (Balasuriya et al., 2002); primers
([forward: 5’-GGC-GAC-AGC-CTA-CAA-GCT-ACA-3’, reverse: 5’-CGG-CAT-CTG-CAG-TGA-GTG-A-3’] and
probe [5’FAM-TTG-CGG-ACC-CGC-ATC-TGA-CCA-A-TAMRA-3’] and (Westcott et al., 2003); primers
[forward: 5’-GTA-CAC-CGC-AGT-TGG-TAA-CA-3’, reverse: 5’-ACT-TCA-ACA-TGA-CGC-CAC-AC-3’] and
probe [5’FAM-TGG-TTC-ACT-CAC-TGC-AGA-TGC-CGG-TAMRA-3’]). It should be noted, however, that
genomic variation among field isolates of EAV could reduce the sensitivity of both RT-PCR and rRT-PCR
assays, even when the primers and probe are based on the most conserved region of the EAV genome
(ORF 7 [Lu et al., 2007]).
In the absence of general agreement on a complete consensus or universal primer set for EAV, and since
no RT-PCR assay can determine the actual infectivity of a sample, there is a value to performing virus
isolation in conjunction with RT-PCR or rRT-PCR for the identification of virus in clinical or post-mortem
specimens.
Strains of EAV isolated from different regions of the world have been classified into different phylogenetic
groups by sequence analysis of the GP3, GP5 and M envelope protein genes (ORFs 3, 5 and 6,
respectively) and the nucleocapsid (N) protein gene (ORF 7 [Balasuriya et al., 1998; Stadejek et al., 1999]).
The GP5 gene has been found to be most useful and reliable for this purpose (Stadejek et al., 1999). The
relationships between strains demonstrated by nucleotide sequencing are a useful molecular
epidemiological tool for tracing the origin of outbreaks of EVA (Balasuriya et al., 1998; Balasuriya et al.,
2004).
c)
Histopathological and immunohistochemical methods
Where mortality is associated with a suspected outbreak of EVA, a wide range of tissues should be
examined for histological evidence of panvasculitis that is especially pronounced in the small arteries
throughout the body, particularly in the caecum, colon, spleen, associated lymphatic glands and adrenal
cortex (Crawford & Henson, 1973; Del Piero, 2000; Jones et al., 1957). The presence of a disseminated
necrotising arteritis involving endothelial and medial cells of affected vessels is considered to be
pathognomonic of EVA. The characteristic vascular lesions present in the mature animal are not, however, a
prominent a feature in many cases of EAV-related abortion.
EAV antigen can be identified in various tissues of EVA-affected animals either in the presence or absence
of lesions (Del Piero, 2000). Antigen has been demonstrated in lung, heart, liver and spleen and the
placenta of aborted fetuses (Del Piero, 2000). Immunohistochemical examination of biopsied skin specimens
has also been investigated as a means of confirming acute EAV infection. Though of some value, it is not
entirely reliable for the diagnosis of the disease. Viral antigen can be detected within the cytoplasm of
infected cells by immunofluorescence using conjugated equine polyclonal anti-EAV serum (Crawford &
Henson, 1973), or by the ABC technique using mouse MAbs to the GP5 (Glaser et al., 1995) or N proteins of
the virus (Del Piero, 2000).
2.
Serological tests
A variety of serological tests including neutralisation (microneutralisation [Senne et al., 1985] and plaque
reduction [McCollum, 1970] (VN)), the complement fixation (CF) test (Fukunaga & McCollum, 1977), the indirect
fluorescent antibody test (Crawford & Henson, 1973), the agar gel immunodiffusion (Crawford & Henson, 1973),
the ELISA (Cho et al., 2000; Hedges et al., 1998; Kondo et al., 1998; Nugent et al., 2000) and the fluorescent
microsphere immunoassay (MIA) (Go, pers. comm.) have been used to detect antibody to EAV.
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Chapter 2.5.10. — Equine viral arteritis
The test currently in widest international use to diagnose infection, carry out seroprevalence studies, and test
horses for export, is a microneutralisation test in the presence of complement. It has also been used to screen
fetal heart blood for the retrospective diagnosis of cases of EVA-related abortion. Apart from the VN test, the CF
test has been used for diagnosing recent EAV infection as complement-fixing antibodies are relatively short-lived
in duration (Fukunaga & McCollum, 1977). In contrast, neutralising antibody titres to EAV frequently persist for
several years after natural infection (Timoney & McCollum, 1993). Although a number of ELISAs have been
developed (Cho et al., 2000; Hedges et al., 1998; Kondo et al., 1998; Nugent et al., 2000), none has as yet been
as extensively validated as the VN, though some appear to offer nearly comparable sensitivity and specificity
(Hedges et al., 1998; Nugent et al., 2000). Unlike the VN test, a positive reaction in the ELISA is not necessarily
reflective of the protective immune status of an individual horse to EAV as both non-neutralising and neutralising
antibodies are involved.
Antiserum to unpurified EAV has been prepared in horses and in rabbits using conventional immunisation
protocols. Also, mouse monoclonal and monospecific rabbit antibodies have been developed to the nucleocapsid
protein (N) major envelope glycoprotein (GP5), and unglycosylated envelope protein (M) of EAV (Balasuriya et
al., 1997; Glaser et al., 1995).
OIE Standard Sera for EAV are available2 and these can facilitate international standardisation of the
microneutralisation test and ELISA.
Only one major serotype of EAV has been recognised so far (McCollum, 1970; Timoney & McCollum, 1993). This
is represented by the prototype Bucyrus strain (ATCC VR 796). The reference virus used in the EAV VN test is
the CVL-Bucyrus (Weybridge) strain. Virus stock is grown in the RK-13 cell line, clarified of cellular debris by lowspeed centrifugation and stored in aliquots at –70°C. Several frozen aliquots are thawed and the infectivity of the
stock virus is determined by titration in RK-13 cells.
a)
Virus neutralisation (a prescribed test for international trade)
The VN test is used to screen stallions for evidence of EAV infection and to determine whether there is a
need to attempt virus detection in semen using cell culture or RT-PCR assay. It is also used for diagnostic
purposes to confirm infection in suspect cases of EVA. The VN test procedure in current widest use is that
developed by the National Veterinary Service Laboratories of the United States Department of Agriculture
(Senne et al., 1985). It is important to obtain a sterile blood sample as bacterial contamination of serum can
interfere with the test result. It is recommended that the test be carried out in RK-13 cells using the approved
CVL-Bucyrus (Weybridge) strain of EAV as reference virus3 (Edwards et al., 1999). Although originally
derived from the prototype Bucyrus virus, the passage history of the CVL (Weybridge) strain is not fully
documented. The sensitivity of the VN test for detection of antibodies to EAV can be significantly influenced
by several factors, especially the source and passage history of the strain of virus used (Edwards et al.,
1999). The CVL-Bucyrus (Weybridge) strain and the highly attenuated MLV vaccine strain of EAV are of
comparable sensitivity for detecting low-titred positive sera, especially from EVA-vaccinated horses. Efforts
are continuing to bring about greater uniformity in the testing protocol and serological results among
laboratories providing the VN or other comparable serological assays for this infection.
2
3

Test procedure
i)
Sera are inactivated for 30 minutes in a water bath at 56°C (control sera, only once).
ii)
Serial twofold dilutions of the inactivated test sera in serum-free cell culture medium (25 µl volumes)
are made in a 96-well, flat-bottomed, cell-culture grade microtitre plate starting at a 1/2 serum dilution
and using duplicate rows of wells for each serum to be tested. Most sera are screened initially at a 1/4
and 1/8 serum dilution (i.e. final serum dilution after addition of an equal volume of the appropriate
dilution of stock virus to each well). Positive samples at the 1/8 dilution can, if desired, be retested and
titrated out for end-point determination. Individual serum controls, together with negative and known
low- and high-titred positive control sera must also be included in each test.
iii)
A dilution of stock virus to contain from 100 to 300 TCID50 (50% tissue culture infective dose) per 25 µl
is prepared using as diluent, serum-free cell culture medium containing antibiotics and fresh guinea-pig
or rabbit complement at a final concentration of 10%.
iv)
25 µl of the appropriate dilution of stock virus is added to every well containing 25 µl of each serum
dilution, except the test serum control wells.
OIE Reference Laboratory for Equine viral arteritis at the Maxwell H. Gluck Equine Research Center, University of
Kentucky, United States of America.
Available from the OIE Reference Laboratory for Equine viral arteritis at the Animal Health Veterinary Laboratories
Agency, Weybridge, United Kingdom.
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Chapter 2.5.10. — Equine viral arteritis
v)
A virus back titration of the working dilution of stock virus is included, using four wells per tenfold
dilution, to confirm the validity of the test results.
vi)
The plates are covered and shaken gently to facilitate mixing of the serum/virus mixtures.
vii)
The plates are incubated for 1 hour at 37°C in a humid atmosphere of 5% CO2 in air.
viii) A suspension of cells from 3- to 5-day-old cultures of RK-13 cells are prepared using a concentration
that will ensure confluent monolayers in the microtitre plate wells within 18–24 hours after seeding.
ix)
100 µl of cell suspension is added to every well, the plates covered with plate lids or sealed with tape
and shaken gently.
x)
The plates are incubated at 37°C in a humid atmosphere of 5% CO2 in air.
xi)
The plates are read microscopically for nonviral CPE after 12–18 hours and again for viral CPE after
48–72 hours’ incubation. The validity of the test is confirmed by establishing that the working dilution of
stock virus contained 30–300 TCID50 virus and that the titres of the positive serum controls are within
0.3 log10 units of their predetermined titres.
A serum dilution is considered to be positive if there is an estimated 75% or preferably a 100% reduction in
the amount of viral CPE in the serum test wells compared with that present in the wells of the lowest virus
control dilution. End-points are then calculated using the Spearman–Kärber method. A titre of 1/4 or greater
is considered to be positive. A negative serum should only have a trace (less than 25%) or no virus
neutralisation at the lowest dilution tested. Antibody titres may, on occasion, be difficult to define as partial
neutralisation may be observed over a range of several serum dilutions. Infrequently, sera will be
encountered that cause toxic changes in the lower dilutions tested. In such cases it may not be possible to
establish whether the sample is negative or a low-titred positive. The problem may be overcome by testing
another serum sample from the animal in question or by retesting the toxic sample using microtitre plates
with confluent monolayers of RK-13 cells that had been seeded the previous day. It has been reported that
the toxicity of the serum can, in some cases, be reduced or eliminated if the sample is adsorbed with packed
RK-13 cells prior to testing or by substituting rabbit in place of guinea-pig complement in the virus diluent.
Vaccination status for equine herpesviruses should be considered when evaluating sera causing non-viral
cytotoxicity. One of the equine herpesvirus vaccines currently available in Europe has been shown to
stimulate antibodies to rabbit kidney cells used in the vaccine production. These, in turn, can give rise to
cytotoxicity, usually in the 1/4 and/or 1/8 serum dilutions and cause difficulties in interpretation of the test
results (Newton et al., 2004).
b)
Enzyme-linked immunosorbent assay
A number of direct or indirect ELISAs have been developed for the detection of antibodies to EAV (Cho et
al., 2000; Hedges et al., 1998; Kondo et al., 1998; Nugent et al., 2000). These have been based on the use
of purified virus or recombinant-derived viral antigens. The usefulness of earlier assays was compromised
by the frequency of false-positive reactions. The latter were associated with the presence of antibodies to
various tissue culture antigens in the sera of horses that had been vaccinated with tissue-culture-derived
antigens. Identification of the importance of the viral GP5 protein in stimulation of the humoral antibody
response to EAV led to the development of several ELISAs that employ a portion of, or the entire
recombinant protein produced in a bacterial or baculovirus expression system (Cho et al., 2000; Hedges et
al., 1998). Most recently, an ovalbumin-conjugated synthetic peptide representing amino acids 81–106 of the
GP5 protein has been used (Nugent et al., 2000). Some of these assays appear to offer nearly comparable
sensitivity and specificity to the VN test and may detect EAV-specific antibodies prior to a positive reaction
being obtainable in the VN test. False-negative reactions can occur, however, with some of these assays.
Screening a random peptide-phage library with polyclonal sera from EAV-infected horses led to the
identification of ligands, which were purified and used as antigen in an ELISA for EAV. No correlation was
found, however, between absorbency values obtained with this assay and neutralising antibody titres,
indicating that the antibodies being detected were largely against nonsurface epitopes of the virus. An ELISA
based on the use of a combination of the GP5, M or N structural proteins of EAV expressed from
recombinant baculoviruses successfully detected viral antibody in naturally or experimentally infected horses
but not in EVA-vaccinated animals (Hedges et al., 1998). Of major importance with respect to any GP5
protein-based ELISA for EAV is the fact that test sensitivity will vary depending on the ectodomain
sequence(s) of this viral protein used in the assay. Considerable amino acid sequence variation within this
domain has been found between isolates of EAV. To maximise sensitivity of a GP5-based ELISA, it may be
necessary to include multiple ectodomain sequences representative of known phenotypically different
isolates of EAV rather than depend on a single ectodomain sequence. Two more recently described ELISAs
appear to offer most promise as reliable serodiagnostic tests for EAV infection (Cho et al., 2000; Nugent et
al., 2000). A blocking ELISA involving MAbs produced against the GP5 protein was reported to have a
sensitivity of 99.4% and a specificity of 97.7% compared with the VN test (Cho et al., 2000). Another assay,
a GP5 ovalbumin-conjugated synthetic peptide ELISA was shown to have a sensitivity and specificity of
96.75% and 95.6%, respectively, using a panel of 400 VN positive sera and 400 VN negative samples
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Chapter 2.5.10. — Equine viral arteritis
(Nugent et al., 2000). It is expected that an ELISA will soon be available, having very similar if not equivalent
sensitivity and specificity to the VN test.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
A number of experimental and commercial vaccines have been developed against EVA. Currently, there are two
commercially available vaccines, both tissue-culture derived. The first is a modified live virus (MLV) vaccine
prepared from virus that has been attenuated for horses by multiple serial transfers in equine and rabbit cell
cultures (Doll et al., 1968; McCollum, 1970). This vaccine is licensed for use in stallions, nonpregnant mares and
in nonbreeding horses. Whereas nonbreeding horses can be vaccinated at any time, stallions and mares should
be vaccinated not less than 3 weeks prior to breeding. The vaccine is not recommended for use in pregnant
mares, especially in the last 2 months of gestation, nor in foals under 6 weeks of age unless in the face of
significant risk of exposure to natural infection. The vaccine is commercially available in the USA and Canada. It
has also been used in New Zealand, subject to ministerial controls, to aid in that country’s EVA eradication
programme.
The second commercially available vaccine against EVA is an inactivated product prepared from virus grown in
equine cell culture, which is filtered, chemically inactivated and then combined with a metabolisable adjuvant. This
vaccine is licensed for use in nonbreeding and breeding horses. In the absence of appropriate safety data, the
vaccine is currently not recommended for use in pregnant mares. The initial vaccination regimen involves two
doses of vaccine administered intramuscularly 3–6 weeks apart. Booster vaccination at 6-month intervals is
recommended by the manufacturer. The inactivated vaccine is licensed for commercial use in certain European
countries, including Denmark, France, Germany, Hungary, Ireland, Sweden and the United Kingdom.
An additional inactivated vaccine against EVA has been developed in Japan for use should an outbreak of EVA
occur in that country. It is an aqueous formalin-inactivated vaccine that has been shown to be safe and effective
for use in nonbreeding and breeding horses. For optimal immunisation with this vaccine, horses require a primary
course of two injections given at an interval of 4 weeks, with a booster dose administered every 6–12 months. As
the vaccine is currently not commercially available, no details can be provided on its production.
Guidelines for the production of veterinary vaccines are given in Chapter 1.1.6 Principles of veterinary vaccine
production. The guidelines given here and in chapter 1.1.6 are intended to be general in nature and may be
supplemented by national and regional requirements.
1.
Seed management
a)
Characteristics of the seed
Both MLV and inactivated commercial vaccines are derived from the prototype Bucyrus strain of EAV (ATCC
VR 796). Available evidence points to the existence of only one major serotype of the virus, and strain
variation is not considered to be of significance in relation to vaccine efficacy (McCollum, 1970; Timoney &
McCollum, 1993).
In the case of the MLV vaccine, the prototype virus was attenuated by serial passage in primary cultures of
horse kidney (HK-131), rabbit kidney (RK-111), and a diploid equine dermal cell line, ATCC CCL57 (ECID24) (Doll et al., 1968; McCollum, 1970). The indications from the use of this vaccine are that the virus is safe
and immunogenic between its 80th and 111th passage in primary rabbit kidney (Doll et al., 1968; McCollum,
1970; Timoney et al., 1988).
The inactivated adjuvanted vaccine is prepared from the unattenuated prototype Bucyrus strain of EAV
(ATCC VR 796) that has been plaque purified and in its fourth serial passage in the diploid equine dermal
cell line (ECID-4). After growth in cell culture, the virus is then purified by filtration before being chemically
inactivated and adjuvanted.
Suitable lots of master seed virus for each vaccine should be maintained in liquid nitrogen or its equivalent.
b)
Method of culture
The virus for both MLV and inactivated vaccines should be grown in a stable cell culture system, such as
equine dermal cells, using an appropriate medium supplemented with sterile bovine serum or bovine serum
albumin as replacement for bovine serum in the growth medium. Cell monolayers should be washed prior to
virus inoculation to remove traces of bovine serum. Extensive virus growth as evidenced by the appearance
of cytopathic changes in 80–100% of the cells should be obtained within 2–3 days.
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c)
Validation as a vaccine
In the case of both MLV and inactivated vaccines, the respective virus strains should be grown in an
appropriate cell culture system that has been officially approved for vaccine production and confirmed to be
free from extraneous bacteria, fungi, mycoplasmas and viruses (Moore, 1986). The identity of the vaccine
virus in the master seed should be confirmed by neutralisation with homologous anti-EAV serum. Incomplete
neutralisation of EAV by homologous horse or rabbit antisera has been scientifically documented (Moore,
1986; Senne et al., 1985) and is a problem when screening master seed virus for extraneous viruses and
when attempting to confirm the identity of the vaccine virus. The problem has been circumvented by
reducing the infectivity titre of the master seed virus below that required for seed virus production before
conducting a neutralisation test on the diluted virus. Virus/serum mixtures are tested for residual live virus by
serial passage in cell culture. No evidence of cytopathic viruses, haemadsorbing viruses, or noncytopathic
strains of bovine virus diarrhoea virus should be found, based on attempted virus isolation in cell culture. If
cells of equine origin are used, they should be confirmed to be free from equine infectious anaemia virus.
The newer technologies of PCR and antigen-capture ELISA may be used as adjuncts to virus isolation in
screening for adventitious agents.
The MLV vaccine has been shown to be both safe and effective for use in stallions and nonpregnant mares
(Timoney et al., 1988). Although not recommended by the manufacturer for pregnant mares, especially in
the last 2 months of gestation, the vaccine has been used to immunise pregnant mares in the face of high
risk of natural exposure to EAV, with minimal, if any, reported adverse effects. Vaccination confers a high
level of protective immunity that persists for at least several years (McCollum, 1970; Timoney & McCollum,
1993). Based on experimental studies and extensive field use of the vaccine since 1985, there is no
evidence of back reversion to virulence of the vaccine virus, nor of recombination of the vaccine virus with
naturally occurring strains of EAV. Furthermore, there is no confirmed evidence that the attenuated strain of
EAV in the current vaccine localises and sets up the carrier state in the reproductive tract of the vaccinated
stallion (Timoney & McCollum, 1993; 2000; Timoney et al., 1988).
The commercial inactivated vaccine has been shown to be nonreactive and safe for use in healthy
nonbreeding and breeding horses. Transient local reactions may be observed in less than 10% of horses
vaccinated with the inactivated vaccine. Limited field studies of this vaccine indicate that it is immunogenic,
stimulating a satisfactory degree of immunity, the duration of which has yet to be reported.
Although there are no published reports on the efficacy of either commercial vaccine in preventing
establishment of the carrier state in the stallion, an experimental aqueous formalin inactivated vaccine
against EVA has been shown to prevent virus persistence in the reproductive tract of vaccinated stallions
following subsequent challenge with EAV (Fukunaga et al., 1992).
2.
Method of manufacture
Both the MLV and inactivated vaccines are produced by cultivation of the respective seed viruses in an equine
dermal cell system. Cell monolayers should be washed prior to inoculation with seed virus to remove traces of
bovine serum in the growth medium. Inoculated cultures should be maintained on an appropriate maintenance
medium. Harvesting of infected cultures should take place when almost the entire cell sheet shows the
characteristic CPE. Supernatant fluid and cells are harvested and clarified of cellular debris and unwanted
material by filtration. In the case of the inactivated vaccine, the purified virus is then chemically inactivated and
adjuvanted with a metabolisable adjuvant.
3.
In-process control
The MLV and inactivated vaccines should be produced in a stable cell line that has been tested for identity and
confirmed to be free from contamination by bacteria, fungi, mycoplasmas or other adventitious agents. In addition
to the preproduction testing of the master seed virus for each vaccine and the cell line for adventitious
contaminants, the cell cultures infected with the respective vaccine viruses should be examined macroscopically
for evidence of microbial growth or other extraneous contamination during the incubation period. If growth in a
culture vessel cannot be reliably determined by visual examination, subculture, microscopic examination, or both
should be carried out.
4.
Batch control
a)
Sterility
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
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b)
Safety
In the case of both MLV and inactivated vaccines, each production lot of vaccine should be checked for
extraneous bacterial, fungal and mycoplasmal contaminants. The vaccine should be safety tested by the
intramuscular inoculation of at least two horses seronegative for neutralising antibodies to EAV with one
vaccine dose of lyophilised virus each (Moore, 1986). None of the inoculated horses should develop any
clinical signs of disease other than mild pyrexia during the ensuing 2-week observation period. In addition,
nasopharyngeal swabs should be collected daily from each horse for attempted virus isolation; white blood
cell counts and body temperatures should also be determined on a daily basis. No significant febrile or
haematological changes should supervene following vaccination (Timoney & McCollum, 1993; Timoney et
al., 1988). Limited shedding of vaccine virus by the respiratory route for at most 7 days may be
demonstrated in the occasional vaccinated horse (Timoney et al., 1988). There is no evidence of persistence
of the vaccine virus in the reproductive tract after vaccination of stallions (Timoney & McCollum, 1993; 2000;
Timoney et al., 1988).
To ensure complete inactivation of the vaccine virus, each serial lot of the inactivated vaccine should be
checked for viable virus by three serial passages in equine dermal cells and by direct fluorescent antibody
staining with specific EAV conjugate before being combined with adjuvant. This should be followed by a
safety test in guinea-pigs and mice.
c)
Potency
Potency of the vaccine in the final containers is determined by plaque infectivity assay in monolayer cultures
of equine dermal cells and by a vaccination challenge test in horses (Moore, 1986). The vaccine must be
tested in triplicate in cell culture, the mean infectivity titre calculated and the dose rate determined on the
basis that each dose of vaccine shall contain not less than 3 × 104 plaque-forming units of attenuated EAV.
The in-vivo potency of the MLV and inactivated vaccines is evaluated in a single vaccination challenge test
using 17–20 vaccinated and 5–7 control horses or in two tests each comprising ten vaccinates and five
controls.
The viral antigen concentration in the inactivated vaccine is over one-thousand times the concentration of
viral antigen present in the MLV vaccine.
d)
Duration of immunity
Detectable neutralising antibody titres to EAV should develop in the majority of horses within 1–2 weeks of
vaccination with the MLV vaccine (Timoney & McCollum, 1993; Timoney et al., 1988). Reported responses
to primary vaccination have been variable in a couple of studies. In one stallion vaccination study, there was
a rapid fall in antibody titres with a significant number of animals reverting to seronegativity 1–3 months after
vaccination (Timoney et al., 1988). On the other hand, other studies have been characterised by an
excellent durable response, with persistence of high VN levels for at least 1–2 years. Revaccination with this
vaccine results in an excellent anamnestic response, with the development of high antibody titres that
remain relatively undiminished for several years (Timoney & McCollum, 1993).
Experimental studies have shown that most horses vaccinated with the inactivated vaccine develop low to
moderate neutralising antibody titres to EAV by day 14 after the second vaccination. There is no published
information on the duration of immunity conferred by this vaccine.
e)
Stability
The lyophilised MLV vaccine can be stored for at least 3–4 years at 2–7°C without loss in infectivity,
provided it is kept in the dark. Infectivity is preserved for much longer periods if vaccine is frozen at –20°C or
below. Once rehydrated, however, the vaccine should be used within 1 hour or else destroyed. The
inactivated vaccine is stored as a liquid suspension at 2–8°C, with no loss of potency for at least 1 year,
provided it is protected from light.
f)
Preservatives
The preservatives added to the MLV and inactivated vaccines are neomycin, polymyxin B and amphotericin B.
g)
Precautions (hazards)
Pregnant mares should not be vaccinated with the MLV vaccine during the last 2 months of gestation, as
there is a risk, albeit minimal, of fetal invasion by the vaccine virus. The possibility of a vaccinally induced
anaphylactic reaction, though very rare, could result from the administration of either the MLV or inactivated
vaccine. In the absence of appropriate safety data, the inactivated vaccine is currently not recommended for
use in pregnant mares.
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5.
Tests on the final product
a)
Safety
With the exception of the inactivated vaccine, which needs to be sterility tested a second time to ensure
freedom from contamination, no further safety tests are required on the inactivated or MLV vaccines.
b)
Potency
No potency tests additional to those conducted on each production lot of the MLV or inactivated vaccines
are required on either final product.
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TIMONEY P.J. & MACLACHLAN N.J. (1998). Serologic and molecular characterization of an abortigenic strain of
equine arteritis virus isolated from infective frozen semen and an aborted equine fetus. J. Am. Vet. Med. Assoc.,
213, 1586–1589.
BALASURIYA U.B., HEDGES J.F., SMALLEY V.L., NAVARETTE A., MCCOLLUM W.H., TIMONEY P.J., SNIJDER E.J. &
MACLACHLAN N.J. (2004). Genetic characterization of equine arteritis virus during persistent infection of stallions. J.
Gen. Virol., 85, 379–390.
BALASURIYA U.B.R., LEUTENEGGER C.M., TOPOL J.B., MCCOLLUM W.H., TIMONEY P.J. & MACLACHLAN N.J. (2002).
Detection of equine arteritis virus by real-time TaqMan® reverse transcription-PCR assay. J. Virol. Methods, 101,
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BALASURIYA U.B.R., PATTON J.F., ROSSITO P.V., TIMONEY P.J., MCCOLLUM W.H. & MACLACHLAN N.J. (1997).
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CAVANAGH D. (1997). Nidovirales: A new order comprising Coronaviridae and Arteriviridae. Arch. Virol., 142, 629–
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CHO H.J., ENTZ S.C., DEREGT D., JORDAN L.T., TIMONEY P.J. & MCCOLLUM W.H. (2000). Detection of antibodies to
equine arteritis virus by a monoclonal antibody-based blocking ELISA. Can. J. Vet. Res., 64, 38–43.
CRAWFORD T.B. & HENSON J.B. (1973). Immunofluorescent, light microscopic and immunologic studies of equine
viral arteritis. Proceedings of the Third International Conference on Equine Infectious Diseases, Paris, 1972.
Karger, Basel, Switzerland, 282–302.
DEL PIERO F. (2000). Equine viral arteritis. Vet. Pathol., 37, 287–296.
DOLL E.R., BRYANS J.T., WILSON J.C. & MCCOLLUM W.H. (1968). Immunisation against equine viral arteritis using
modified live virus propagated in cell cultures of rabbit kidney. Cornell Vet., 48, 497–524.
EDWARDS S., CASTILLO-OLIVARES J., CULLINANE A., LABLE J., LENIHAN P., MUMFORD J.A., PATON D.J., PEARSON J.E.,
SINCLAIR R., WESTCOTT D.G.F., WOOD J.L.N., ZIENTARA S. & NELLY M. (1999). International harmonisation of
laboratory diagnostic tests for equine viral arteritis. Proceedings of the Eighth International Conference on Equine
Infectious Diseases, Dubai, UAE, 1998, 359–362.
FUKUNAGA Y. & MCCOLLUM W.H. (1977). Complement fixation reactions in equine viral arteritis. Am. J. Vet. Res.,
38, 2043–2046.
FUKUNAGA Y., WADA R., MATSUMURA T., ANZAI T., IMAGAWA H., SUGIURA T., KUMANOMIDO T., KANEMARU T. & KAMADA
M. (1992). An attempt to protect against persistent infection of equine viral arteritis in the reproductive tract of
stallions using formalin inactivated-virus vaccine. Proceedings of the Sixth International Conference on Equine
Infectious Diseases, Cambridge, UK, 1991, 239–244.
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FUKUNAGA Y., WADA R., SUGITA S., FUJITA Y., NAMBO Y., IMAGAWA H., KANEMARU T., KAMADA M., KOMATSU N. &
AKASHI H. (2000). In vitro detection of equine arteritis virus from seminal plasma for identification of carrier
stallions. J. Vet. Med. Sci., 62, 643–646.
GILBERT S.A., TIMONEY P.J., MCCOLLUM W.H. & DEREGT D. (1997). Detection of equine arteritis virus in the semen
of carrier stallions using a sensitive nested PCR assay. J. Clin. Microbiol., 35, 2181–2183.
GLASER A.L., DE VRIES A.A.F. & DUBOVI E.J. (1995). Comparison of equine arteritis virus isolates using neutralizing
monoclonal antibodies and identification of sequence changes in GL associated with neutralization resistance. J.
Gen. Virol., 76, 2223–2233.
HEDGES J.F., BALASURIYA U.B.R., SHABBIR A., TIMONEY P.J., MCCOLLUM W.H., YILMA T. & MACLACHLAN N.J. (1998).
Detection of antibodies to equine arteritis virus by enzyme linked immunosorbant assays utilizing GL, M and N
proteins expressed from recombinant baculoviruses. J. Virol. Methods, 76, 127–137.
JONES T.C., DOLL E.R. & BRYANS J.T. (1957). The lesions of equine viral arteritis. Cornell Vet., 47, 52–68.
KONDO T., FUKUNAGA Y., SEKIGUCHI K., SUGIURA T. & IMAGAWA H. (1998). Enzyme-linked immunosorbent assay for
serological survey of equine arteritis virus in racehorses. J. Vet. Med. Sci., 60, 1043–1045.
LITTLE T.V., DEREGT D., MCCOLLUM W.H., & TIMONEY P.J. (1995). Evaluation of an immunocytochemical method for
rapid detection and identification of equine arteritis virus in natural cases of infection. Proceedings of the Seventh
International Conference on Equine Infectious Diseases, Tokyo, Japan, 1994, 27–31.
LU Z., BRANSCUM A., SHUCK K.M., ZANG J., DUBOVI E., TIMONEY P.J. & BALASURIYA U.B.R. (2007). Detection of equine
arteritis virus nucleic acid in equine semen and tissue culture fluid. J. Vet. Diagn. Invest.
MCCOLLUM W.H. (1970). Vaccination for equine viral arteritis. Proceedings of the Second International Conference
on Equine Infectious Diseases, Paris, 1969, Karger, Basle, Switzerland, 143–151.
MCCOLLUM W.H., PRICKETT M.E. & BRYANS J.T. (1971). Temporal distribution of equine arteritis virus in respiratory
mucosa, tissues and body fluids of horses infected by inhalation. Res. Vet. Sci., 2, 459–464.
MOORE B.O. (1986). Development and evaluation of three equine vaccines. Irish Vet. J., 40, 105–107.
NEWTON J.R., GERAGHTY R.J., CASTILLO-OLIVARES J., CARDWELL M. & MUMFORD J.A. (2004). Evidence that use of an
inactivated equine herpesvirus vaccine induces serum cytotoxicity affecting the equine arteritis virus neutralisation
test. Vaccine, 22, 4117–4123.
NUGENT J., SINCLAIR R., DEVRIES A.A.F., EBERHARDT R.Y., CASTILLO-OLIVARES J., DAVIS POYNTER N., ROTTIER P.J.M.
& MUMFORD J.A. (2000). Development and evaluation of ELISA procedures to detect antibodies against the major
envelope protein (GL) of equine arteritis virus. J. Virol. Methods, 90, 167–183.
SENNE D.A., PEARSON J.E. & CABREY E.A. (1985). Equine viral arteritis: A standard procedure for the virus
neutralisation test and comparison of results of a proficiency test performed at five laboratories. Proc. U.S. Anim.
Health Assoc., 89, 29–34.
STADEJEK T., BJORKLUND H., BASCUNANA C.R., CIABATTI I.M., SCICLUNA M.T., AMADDEO D., MCCOLLUM W.H.,
AUTORINO G.L., TIMONEY P.J., PATON D.J., KLINGEBORN B. & BELAK S. (1999). Genetic diversity of equine arteritis
virus. J. Gen. Virol., 80, 691–699.
TIMONEY P.J., BRUSER C.A., MCCOLLUM W.H., HOLYOAK G.R. & LITTLE T.V. (2004). Comparative sensitivity of LLCMK2, RK-13, vero and equine dermis cell lines for primary isolation and propagation of equine arteritis virus. In:
Proceedings of the International Workshop on the Diagnosis of Equine Arteritis Virus Infection, Timoney P.J., ed.
M.H. Cluck Equine Research Center, 20–21 October 2004, Lexington, Kentucky, USA, pp 26–27.
TIMONEY P.J. & MCCOLLUM W.H. (1993). Equine viral arteritis. Vet. Clin. North Am. Equine Pract., 9, 295–309.
TIMONEY P.J. & MCCOLLUM W.H. (2000). Equine viral arteritis: Further characterization of the carrier state in the
stallion. J. Reprod. Fertil. (Suppl.), 56, 3–11.
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TIMONEY P.J., UMPHENOUR N.W. & MCCOLLUM W.H. (1988). Safety evaluation of a commercial modified live equine
arteritis virus vaccine for use in stallions. Proceedings of the Fifth International Conference on Equine Infectious
Diseases, Lexington, 1987, University Press of Kentucky, Lexington, Kentucky, USA, 19–27.
VAALA W.E., HAMIR A.N., DUBOVI E.J., TIMONEY P.J. & RUIZ B. (1992). Fatal congenitally acquired equine arteritis
virus infection in a neonatal foal. Equine Vet. J., 24, 155–158.
WEBER H., BECKMANN K. & HAAS L. (2006). Fallbericht. Equines arteritisvirus (EAV) als aborterreger bei alpacas?
Dtsch. Tierarztl. Wschr., 113, 162–163.
WESTCOTT D.G., KING D.P., DREW T.W., NOWOTNY N., KINDERMANN J., HANNANT D., BELAK S. & PATON D.J. (2003).
Use of an internal standard in a closed one-tube RT-PCR for the detection of equine arteritis virus RNA with
fluorescent probes. Vet. Res., 34, 165–176.
*
* *
NB: There are OIE Reference Laboratories for Equine viral arteritis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for equine viral arteritis
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NB: Version amended following delisting of the mallein test by the World Assembly in May 2012
CHAPTER 2.5.11.
GLANDERS
SUMMARY
Glanders is a contagious and fatal disease of horses, donkeys, and mules, and is caused by
infection with the bacterium Burkholderia mallei (the name recently changed from Pseudomonas
mallei and was previously classified as Pfeifferella, Loefflerella, Malleomyces or Actinobacillus).
The disease causes nodules and ulcerations in the upper respiratory tract and lungs. A skin form
also occurs, known as ‘farcy’. Control of glanders requires testing of suspect clinical cases,
screening of apparently normal equids, and elimination of positive reactors. It is transmitted to
humans and all infected/contaminated or potentially infected/contaminated material must be
handled in a laboratory that meets the requirements for Containment Group 3 pathogens.
Identification of the agent: Smears from fresh material may reveal Gram-negative nonsporulating,
nonencapsulated rods. The presence of a capsule-like cover has been demonstrated by electron
microscopy. The bacteria grow aerobically and prefer media that contain glycerol. Unlike the
Pseudomonas species and the closely related bacterium Burkholderia pseudomallei, Burkholderia
mallei is nonmotile. Guinea-pigs are highly susceptible, and males can be used, if strictly
necessary, to recover the organism from a heavily contaminated sample. Commercially available
biochemical identification kits lack diagnostic sensitivity. Specific monoclonal antibodies and
polymerase chain reaction (PCR) as well as real-time PCR assays are available. The latter have
also been evaluated in recent outbreaks.
Mallein and serological tests: Complement fixation test and enzyme-linked immunosorbent
assays are the most accurate and reliable serological tests for diagnostic use. The mallein test is a
sensitive and specific clinical test for hypersensitivity against Burkholderia mallei. Mallein, a water
soluble protein fraction of the organism, is injected subcutaneously, intradermo-palpebrally or given
by eyedrop. In infected animals, the skin or the eyelid swells markedly within 1–2 days. A rose
bengal plate agglutination test has recently been developed in Russia; it has been validated in
Russia only.
Requirements for vaccines and diagnostic biologicals: There are no vaccines. Mallein purified
protein derivative is currently available commercially from the Central Veterinary Control and
Research Institute, 06020 Etlik, Ankara, Turkey.
A. INTRODUCTION
Glanders is a bacterial disease of perissodactyls or odd-toed ungulates with zoonotic potential known since
ancient times. It is caused by the bacterium Burkholderia mallei (the name recently changed from Pseudomonas
mallei (Yabuuchi et al., 1992) and has been classified in the past as Pfeifferella, Loefflerella, Malleomyces or
Actinobacillus). Outbreaks of the disease may occur in felines living in the wild or in zoological gardens.
Susceptibility to glanders has been proved in camels, bears, wolves and dogs. Carnivores may become infected
by eating infected meat, but cattle and pigs are resistant (Minett, 1959). Small ruminants may also be infected if
kept in close contact to glanderous horses (Wittig et al., 2006). Glanders in the acute form occurs most frequently
in donkeys and mules with high fever and respiratory signs (swollen nostrils, dyspnoea, and pneumonia); death
occurs within a few days. In horses, glanders generally takes a more chronic course and they may survive for
several years. Chronic and subclinical ‘occult’ cases are dangerous sources of infection due to permanent or
intermittent shedding of bacteria (Wittig et al., 2006).
In horses, inflammatory nodules and ulcers develop in the nasal passages and give rise to a sticky yellow
discharge, accompanied by enlarged firm submaxillary lymph nodes. Stellate scarring follows upon healing of the
ulcers. The formation of nodular abscesses in the lungs is accompanied by progressive debility, febrile episodes,
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Chapter 2.5.11. — Glanders
coughing and dyspnoea. Diarrhoea and polyuria can also occur. In the skin form (‘farcy’), the lymphatics are
enlarged and nodular abscesses (‘buds’) of 0.5–2.5 cm develop, which ulcerate and discharge yellow oily pus.
Nodules are regularly found in the liver and spleen. Discharges from the respiratory tract and skin are infective,
and transmission between animals, which is facilitated by close contact, by inhalation, ingestion of contaminated
material (e.g. from infected feed and water troughs), or by inoculation (e.g. via a harness) is common. The
incubation period can range from a few days to many months (Monlux, 1984; Wittig et al., 2006).
Glanders is transmissible to humans by direct contact with diseased animals or infected/contaminated material. In
the untreated acute disease. 95% mortality can occur within 3 weeks (Neubauer et al., 1997). However, survival is
possible if the infected person is treated early and aggressively with multiple systemic antibiotic therapies (Kahn &
Ashford, 2001; Srinivasan et al., 2001). A chronic form with abscessation also occurs (Neubauer et al., 1997).
When handling suspect or known infected animals or fomites, stringent precautions should be taken to prevent
self-infection or transmission of the bacterium to other equids. Laboratory samples should be securely packaged,
kept cool and shipped as outlined in Chapter 1.1.1 Collection and shipment of diagnostic specimens. All
manipulations with potentially infected/contaminated material must be performed in a laboratory that meets the
requirements for Containment Group 3 pathogens as outlined in Chapter 1.1.3 Biosafety and biosecurity in the
veterinary microbiology laboratory and animal facilities.
Glanders has been eradicated from many countries by statutory testing, elimination of infected animals, and
import restrictions. It persists in some Asian, African and South American countries. It can be considered a reemerging disease and may be imported by pet or racing equids into glanders free areas (Neubauer et al., 2005).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
Cases for specific glanders investigation should be differentiated on clinical grounds from other chronic infections
of the nasal mucosae or sinuses, and from strangles (Streptococcus equi infection), ulcerative lymphangitis
(Corynebacterium pseudotuberculosis), pseudotuberculosis (Yersinia pseudotuberculosis) and sporotrichosis
(Sporotrichium spp.). Glanders should be excluded positively from suspected cases of epizootic lymphangitis
(caused by Histoplasma farciminosum), with which it has many clinical similarities. In humans in particular,
glanders should be distinguished from melioidosis (B. pseudomallei infection), which is caused by an organism
with close similarities to B. mallei (Minett, 1959).
a)
Morphology of Burkholderia mallei
The organisms are fairly numerous in smears from fresh lesions, but in older lesions they are scanty (Wilson
& Miles, 1964). They should be stained by methylene blue or Gram stain. The organisms are mainly
extracellular, fairly straight Gram-negative rods with rounded ends, 2–5 µm long and 0.3–0.8 µm wide with
granular inclusions of various size. They often stain irregularly and do not have a readily visible capsule,
under the light microscope, or form spores. The presence of a capsule-like cover has been established by
electron microscopy. This capsule is composed of neutral carbohydrates and serves to protect the cell from
unfavourable environmental factors. Unlike other organisms in the Pseudomonas group and its close relative
Burkholderia pseudomallei, Burkholderia mallei has no flagellae and are therefore nonmotile (Krieg & Holt,
1984; Sprague & Neubauer, 2004). The organisms are difficult to demonstrate in tissue sections, where they
may have a beaded appearance (Miller et al., 1948). In culture media, they vary in appearance depending
on the age of the culture and type of medium. In older cultures, there is much pleomorphism. Branching
filaments form on the surface of broth cultures (Neubauer et al., 2005).
b)
Cultural characteristics
It is preferable to attempt isolation from unopened uncontaminated lesions (Miller et al., 1948). The organism
is aerobic and facultatively anaerobic only in the presence of nitrate (Gilardi, 1985; Krieg & Holt, 1984),
growing optimally at 37°C (Mahaderan et al., 1987). It grows well, but slowly, on ordinary culture media, 72hour incubation of cultures is recommended; glycerol enrichment is particularly useful. After a few days on
glycerol agar, there is a confluent, slightly cream-coloured growth that is smooth, moist, and viscuous. With
continued incubation, the growth thickens and becomes dark brown and tough. It also grows well on glycerol
potato agar and in glycerol broth, on which a slimy pellicle forms. On plain nutrient agar, the growth is much
less luxuriant, and growth is poor on gelatin (Steele, 1980). In samples not obtained under sterile conditions
B. mallei is regularly overgrown by other bacteria.
Alterations to characteristics may occur in vitro, so fresh isolates should be used for identification reactions.
Litmus milk is slightly acidified by B. mallei, and coagulation may occur after long incubation. The organism
reduces nitrates. Although some workers have claimed that glucose is the only carbohydrate that is
fermented (slowly and inconstantly), other workers have shown that if an appropriate medium and indicator
are used, glucose and other carbohydrates, such as arabinose, fructose, galactose and mannose, are
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Chapter 2.5.11. — Glanders
consistently fermented by B. mallei (Evans, 1966). Indole is not produced, horse blood is not haemolysed
and no diffusible pigments are produced in cultures (Krieg & Holt, 1984). A commercial laboratory test kit
(e.g. API [Analytical Profile Index] system: Analytab Products, BioMerieux or Biolog [Hayward, California])
can be used for easy confirmation that an organism belongs to the Pseudomonas group. In general,
commercially available systems are not suited to unambiguously identifying members of the steadily growing
number of species of the genus Burkholderia (Glass & Popovic, 2005). Lack of motility is then of special
relevance. A bacteriophage specific for B. mallei is available (Woods et al., 2002).
All culture media prepared should be subjected to quality control and must support growth of the suspect
organism from a small inoculum. The reference strain should be cultured in parallel with the suspicious
samples to ensure that the tests are working correctly.
In contaminated samples, supplementation of media with substances that inhibit the growth of Gram-positive
organisms (e.g. crystal violet, proflavine) has proven to be of use, as has pretreatment with penicillin
(1000 units/ml for 3 hours at 37°C) (Minett, 1959). A selective medium has been developed (Xie et al., 1980)
composed of polymyxin E (1000 units), bacitracin (250 units), and actidione (0.25 mg) incorporated into
nutrient agar (100 ml) containing glycerine (4%), donkey or horse serum (10%), and ovine haemoglobin or
tryptone agar (0.1%).
Outside the body, the organism has little resistance to drying, heat, light or chemicals, so that survival
beyond 2 weeks is unlikely (Neubauer et al., 1997). Under favourable conditions, however, it can probably
survive a few months. Burkholderia mallei can remain viable in tap water for at least 1 month (Steele, 1980).
For disinfection, benzalkonium chloride or ‘roccal’ (1/2000), sodium hypochlorite (500 ppm available
chlorine), iodine, mercuric chloride in alcohol, and potassium permanganate have been shown to be highly
effective against B. mallei (Mahaderan et al., 1987). Phenolic disinfectants are less effective.
c)
Laboratory animal inoculation
Guinea-pigs, hamsters and cats have been used for diagnosis when necessary. If isolation in a laboratory
animal is considered unavoidable, suspected material is inoculated intraperitoneally into a male guinea-pig.
As this technique has a sensitivity of only 20%, the inoculation of at least five animals is recommended
(Neubauer et al., 1997). Positive material will cause a severe localised peritonitis and orchitis (the Strauss
reaction). The number of organisms and their virulence determines the severity of the lesions. Additional
steps are used when the test material is heavily contaminated (Gould, 1950). The Strauss reaction is not
specific for glanders, and other organisms can elicit it. Bacteriological examination of infected testes should
confirm the specificity of the response obtained.
d)
Confirmation by polymerase chain reaction and real-time PCR
In the past few years, several PCR and real-time PCR assays for the identification of Burkholderia mallei
have been developed (Bauernfeind et al., 1998; Lee et al., 2005; Sprague et al., 2002; Thibault et al., 2004;
Ulrich et al., 2006; U’Ren et al., 2005), but only a PCR and a real-time PCR assay were evaluated using
samples from a recent outbreak of glanders in horses (Scholz et al., 2006; Tomaso et al., 2006). These two
assays will be described in more detail here. However, the robustness of these assays will have to be
demonstrated in the future by interlaboratory studies. The guidelines and precautions outlined in Chapter
1.1.5 Principles and methods of validation of diagnostic assays for infectious diseases have to be taken into
account.
•
DNA preparation
Single colonies are transferred from an agar plate to 200 µl lysis buffer (5× buffer D [PCR Optimation Kit,
Invitrogen, DeShelp, The Netherlands, 1/5 diluted in ultra-pure water]; 0.5% Tween 20 [ICI, American
Limited, Merck, Hohenbrunn, Germany]; 2 mg/ml proteinase K [Roche Diagnostics, Mannheim, Germany]).
After incubation at 56°C for 1 hour and inactivation for 10 minutes at 95°C, 2 and 4 µl of the cleared lysate
are used as template in the PCR or the real-time PCR assay, respectively.
Tissue samples of horses (skin, liver, spleen, lung, and conchae) inactivated and preserved in formalin
(48 hours, 10% v/v) are cut with a scalpel into pieces of 0.5 × 0.5 cm (approximately 500 mg). The
specimens are washed twice in deionised water (10 ml), incubated over night in sterile saline at 4°C, and
minced using liquid nitrogen, a mortar and a pestle. Total DNA is prepared from 50 mg tissue using the
TM
QIAamp Tissue Kit according to the manufacturer’s instructions (Qiagen, Hilden, Germany). DNA is eluted
with 80 µl dH2O of which 4 µl are used as template.
•
PCR assay (Scholz et al., 2006)
The assay may have to be adapted to the PCR instrument used with minor modifications to the cycle
conditions and the concentration of the chemicals used.
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Chapter 2.5.11. — Glanders
The oligonucleotides used in by Scholz et al. (2006) are designed based on differences of the fliP
sequences from B. mallei ATCC 23344T (accession numbers NC_006350, NC_006351) and B. pseudomallei K96243 (accession numbers NC_006348, NC_006349). Primers Bma-IS407-flip-f (5’-TCA-GGT-TTGTAT-GTC-GCT-CGG-3’) and Bma- IS407-flip-r (5’-CTA-GGT-GAA-GCT-CTG-CGC-GAG-3’) are used to
amplify a 989 bp fragment. PCR is done using 50 µl ready-to-go mastermix (Eppendorf, Hamburg,
Germany) and 15 pmol of each primer. Thermal cycling conditions are 94°C for 30 seconds; 65°C for
30 seconds and 72°C for 60 seconds. This cycle is repeated for 35 times. A final elongation step is added
(72°C for 7 minutes). Visualisation of the products is done by UV light after agarose gel (1% w/v in TAE
buffer) electrophoresis and staining with ethidium bromide. No template controls containing PCR-grade
water instead of template and positive controls containing DNA of B. mallei have to be included in each run
to detect contamination by amplicons of former runs or amplification failure.
The lower detection limit is 10 fg or 2 genome equivalents.
•
Real-time PCR assay (Tomaso et al., 2006)
The assay has to be adapted to the real-time PCR instrument used with minor modifications, e.g. the cycling
vials have to be chosen according to the manufacturer’s recommendations and the concentration of the
oligonucleotides may have to be doubled or the labelling of the probes has to be changed. The authors used
TM
real-time PCR system (Stratagene, Amsterdam, The Netherlands) and 96-well plates
a MX3000P
(ThermoFast 96 ABGeneTM, Rapidozym, Berlin, Germany).
The oligonucleotides used by Tomaso et al. (2006) were designed based on differences of the fliP
sequences from B. mallei ATCC 23344T (accession numbers NC_006350, NC_006351) and B. pseudomallei K96243 (accession numbers NC_006348, NC_006349). The fluorogenic probe is synthesised with 6carboxy-fluorescein (FAM) at the 5’-end and black hole quencher 1 (BHQ1) at the 3’-end. Oligonucleotides
used were Bma-flip-f (5’-CCC-ATT-GGC-CCT-ATC-GAA-G-3’), Bma-flip-r (5’-GCC-CGA-CGA-GCA-CCTGAT-T-3’) and probe Bma-flip (5’-6FAM-CAG-GTC-AAC-GAG-CTT-CAC-GCG-GAT-C-BHQ1-3’).
The 25 µl reaction mixture consists of 12.5 µl 2× TaqManTM Universal MasterMix (Applied Biosystems,
Foster City, USA), 0.1 µl of each primer (10 pmol/µl), 0.1 µl of the probe (10 pmol/µl) and 4 µl sample.
Thermal cycling conditions are 50°C for 2 minutes; 95°C for 10 minutes; 45 (50) cycles at 95°C for
25 seconds and 63°C for 1 minute. Possible contaminations with amplification products from former
reactions are inactivated by an initial incubation step using uracil N’-glycosilase.
The authors suggest to include an internal inhibition control based on a bacteriophage lambda gene target
(Lambda-F [5’-ATG-CCA-CGT-AAG-CGA-AAC-A-3] Lambda-R [5’-GCA-TAA-ACG-AAG-CAG-TCG-AGT-3’],
Lam-YAK [5’-YAK-ACC-TTA-CCG-AAA-TCG-GTA-CGG-ATA-CCG-C-DB-3’]), which can be titrated to give
reproducible cycle threshold values. However, depending on the sample material a house keeping gene
targeting PCR may be used additionally or as an alternative. No template controls containing 4 µl of PCRgrade water instead and positive controls containing DNA of B. mallei have to be included in each run to
detect amplicon contamination or amplification failure.
The linear range of the assay was determined to cover concentrations from 240 pg to 70 fg bacterial
DNA/reaction. The lower limit of detection defined as the lowest amount of DNA that was consistently
detectable in three runs with eight measurements each is 60 fg DNA or four genome equivalents (95%
probability). The intra-assay variability of the fliP PCR assay for 35 pg DNA/reaction is 0.68 % (based on Ct
values) and for 875 fg 1.34%, respectively. The inter-assay variability for 35 pg DNA/reaction is 0.89%
(based on Ct values) and for 875 fg DNA 2.76 %, respectively.
e)
Other methods
The genome of the Burkholderia mallei type strain ATCC 23344T was sequenced in 2004 (Nierman et al.,
2004). Several genomes of other isolates followed and revealed a wide genomic plasticity. Passages in
different host species or culture media may provoke considerable sequence alterations (Romero et al.,
2006). The loss of the ability to produce LPS and/or capsule polysaccharide upon ongoing culture due to
mutation is a well known fact and results in reduced or absent virulence and influences serologic tests
(Neubauer et al., 2005). Several molecular typing techniques have successfully been introduced. Simple
molecular techniques like PCR-restriction fragment length polymorphism (Tanpiboonsak et al., 2004) and
pulsed field gel electrophoresis (Chantratita et al., 2006) can be used for further discrimination of isolates.
Ribotyping using restriction enzymes PstI and EcoRI in combination with an E. coli 18-mer rDNA probe
produced 17 distinct ribotypes within 25 B. mallei isolates (Harvey & Minter, 2005). These techniques are
still the in-house tests of specialised laboratories as an extensive strain collection is necessary. Multilocus
sequence typing (MLST) can be done with purified DNA so there is no need for excessive cultivation of the
agent or the keeping of strain collections. Web-based analysis might even enhance diagnostics (Godoy et
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Chapter 2.5.11. — Glanders
al., 2003). No specific histopathology features can be described for lesions caused by B. mallei. For
immunuhistochemical analysis, B. mallei specific hyperimmune sera of rabbits can be used.
2.
Serological tests and the mallein test
a)
Complement fixation test (a prescribed test for international trade)
Although not as sensitive as the mallein test, the CF test is an accurate serological test that has been used
for glanders diagnosis for many years (Blood & Radostits, 1989). It is reported to be 90–95% accurate,
serum being positive within 1 week of infection and remaining positive in the case of exacerbation of the
chronic process (Steele, 1980). Recently, however, the specificity of CF testing has been questioned
(Neubauer et al., 2005).
•
Antigen preparation (Kelser & Reynolds, 1935)
i)
Flasks of beef infusion broth with 3% glycerol are inoculated with log-phase growth B. mallei and
incubated at 37°C for 8–12 weeks.
ii)
The cultures are inactivated by exposing the flasks to flowing steam (100°C) for 60 minutes.
iii)
The clear supernatant is decanted and filtered. The filtrate is heated again by exposure to live steam
for 75 minutes on 3 consecutive days, and clarified by centrifugation.
iv)
The clarified product is concentrated to one-tenth the original volume by evaporation on a steam or hot
water bath.
v)
Concentrated antigen is bottled in brown-glass bottles to protect from light and stored at 4°C. Antigen
has been shown to be stable for at least 10 years in this concentrated state.
vi)
Lots of antigen are prepared by diluting the concentrated antigen 1/20 with sterile physiological saline
with 0.5% phenol. The diluted antigen is dispensed into brown-glass vials and store at 4°C. The final
working dilution is determined by a block titration. The final working dilution for CF test use is made at
the time the CF test is performed.
The resulting antigen is primarily lipopolysaccharide. An alternative procedure is to use young cultures by
growing the organism on glycerol–agar slopes for 12 hours and washing off with normal saline. A
suspension of the culture is heated for 1 hour at 70°C and the heat-treated bacterial suspension is used as
antigen. The disadvantage of this antigen preparation method is that the antigen contains all the bacterial
cell components. The antigen should be safety tested by inoculating blood agar plates.
•
Test procedure (NVSL, 2006)
i)
Serum is diluted 1/5 in veronal (barbiturate) buffered saline containing 0.1% gelatin (VBSG) or CFD
(complement fixation diluent – available as tablets) without gelatine.
ii)
Diluted serum is inactivated for 30 minutes at 56°C. The USDA complement fixation protocol calls for
inactivation for 35 minutes (NVSL, 2006). (Serum of equidae other than horses should be inactivated at
63°C for 30 minutes.)
iii)
Twofold dilutions of the sera are prepared in 96-well round-bottom microtitre plates.
iv)
Guinea-pig complement is diluted in the chosen buffer and 5 (or optionally 4) complement haemolytic
units-50% (CH50) are used.
v)
Sera, complement and antigen are reacted in the plates and incubated for 1 hour at 37°C. (An alternate
acceptable procedure is overnight incubation at 4°C.)
vi)
A 2% suspension of sensitised washed sheep red blood cells is added. The USDA protocol calls for
confirmation of positive reactions in a tube test using 3% sheep red blood cells (NVSL, 2006).
vii)
Plates are incubated for 45 minutes at 37°C, and then centrifuged for 5 minutes at 600 g.
A sample that produces 100% haemolysis at the 1/5 dilution is negative, 25–75% haemolysis is suspicious,
and no haemolysis (100% fixation) is positive. Unfortunately, false-positive results can occur, and
B. pseudomallei and B. mallei cross react and cannot be differentiated by serology (Blood & Radostits, 1989;
Neubauer et al., 1997). Also healthy horses can have a false positive CF reaction for a variable period
following a mallein intradermal test.
b)
Enzyme-linked immunosorbent assays
Both plate and membrane (blot) enzyme-linked immunosorbent assays (ELISAs) have been reported for the
serodiagnosis of glanders, but none of these procedures has been shown to differentiate serologically
between B. mallei and B. pseudomallei. Blotting approaches have involved both dipstick dot-blot and
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Chapter 2.5.11. — Glanders
electrophoretically separated and transferred western blot methods (Katz et al., 1999; Verma et al., 1990). A
competitive ELISA that uses an uncharacterised anti-lipopolysaccharide monoclonal antibody has also been
developed and found to be similar to the CF test in performance (Katz et al., 2000). Continuing development
of monoclonal antibody reagents specific for B. mallei antigenic components offers the potential for more
specific ELISAs in the foreseeable future that will help resolve questionable test results of quarantined
imported horses (Burtnick et al., 2002; Feng et al., 2006; Khrapova et al., 1995; Neubauer et al., 1997). At
this time, none of these tests has been validated.
c)
The mallein test
The mallein purified protein derivative (PPD), which is available commercially, is a solution of water-soluble
protein fractions of heat-treated B. mallei. The test depends on infected horses being hypersensitive to
mallein. Advanced clinical cases in horses and acute cases in donkeys and mules may give inconclusive
results requiring additional methods of diagnosis to be employed (Allen, 1929).
•
The intradermo-palpebral test
This is the most sensitive, reliable and specific test for detecting infected perissodactyls or odd-toed
ungulates, and has largely displaced the ophthalmic and subcutaneous tests (Blood & Radostits, 1989):
0.1 ml of concentrated mallein PPD is injected intradermally into the lower eyelid and the test is read at
24 and 48 hours. A positive reaction is characterised by marked oedematous swelling of the eyelid, and
there may be a purulent discharge from the inner canthus or conjunctiva. This is usually accompanied by a
rise in temperature. With a negative response, there is usually no reaction or only a little swelling of the
lower lid.
d)
Other serological tests
The avidin–biotin dot ELISA has been described (Verma et al., 1990), but has not yet been widely used or
validated. The antigen is heat-inactivated bacterial culture that has been concentrated and purified. A dot of
this antigen is placed on a nitrocellulose dipstick that is then used to test for antibody against B. mallei in
equine serum. Using antigen-dotted, preblocked dipsticks, the test can be completed in approximately
1 hour. Serum or whole blood can be used for the test, and partial haemolysis does not impart any
background colour to the antigen-coated area on the nitrocellulose. Recently, polysaccharide microarray
technology has offered a new promising approach to improve sensitivity in serology (Parthasarathy et al.,
2006).
The rose bengal plate agglutination test (RBT) has been described for the diagnosis of glanders in horses
and other susceptible animals; this test has been validated in Russia only. The antigen is a heat-inactivated
bacterial suspension coloured with rose bengal, which is used in a plate agglutination test.
The accuracy of other agglutination tests and precipitin is unsatisfactory for use in control programmes.
Horses with chronic glanders and those in a debilitated condition give negative or inconclusive results.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
No vaccines are available.
Mallein PPD for use in performing the intradermo-palpebral and ophthalmic tests is produced commercially by the
Central Veterinary Control and Research Institute, 06020 Etlik, Ankara, Turkey.
The ID-Lelystad has provided the following information on requirements for mallein PPD.
1.
Seed management
Three strains of Burkholderia mallei are employed in the production of mallein PPD, namely Bogor strain
(originating from Indonesia), Mukteswar strain (India) and Zagreb strain (Yugoslavia). The seed material is kept
as a stock of freeze-dried cultures. The strains are subcultured on to glycerol agar at 37°C for 1–2 days. For
maintaining virulence and antigenicity, the strains may be passaged through guinea-pigs.
2.
Method of manufacture
DorsetHenley medium, enriched by the addition of trace elements, is used for production of mallein PPD. The
liquid medium is inoculated with a thick saline suspension of B. mallei, grown on glycerol agar. The production
medium is incubated at 37°C for about 10 weeks. The bacteria are then killed by steaming for 3 hours in a Koch’s
918
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Chapter 2.5.11. — Glanders
steriliser. The fluid is then passed through a layer of cotton wool to remove coarse bacterial clumps. The resulting
turbid fluid is cleared by membrane filtration, and one part trichloroacetic acid 40% is immediately added to nine
parts culture filtrate. The mixture is allowed to stand overnight and a light brownish to greyish precipitate settles.
The supernatant fluid is pipetted off and discarded. The precipitate is centrifuged for 15 minutes at 2500 g and the
layer of precipitate is washed three or more times in a solution of 5% NaCl, pH 3, until the pH is 2.7. The washed
precipitate is dissolved by stirring with a minimum of an alkaline solvent. The fluid is dark brown and a pH of 6.7
will be obtained. This mallein concentrate has to be centrifuged thoroughly and the supernatant is diluted with an
equal amount of a glucose buffer solution. The protein content of this product is estimated by the Kjeldahl method
and freeze-dried after it has been dispensed into ampoules.
3.
In-process control
During the period of incubation, the flasks are inspected frequently for any signs of contamination, and suspect
flasks are discarded. A typical growth of the B. mallei cultures comprises turbidity, sedimentation, some surface
growth with a tendency towards sinking, and the formation of a conspicuous slightly orange-coloured ring along
the margin of the surface of the medium.
4.
Batch control
Each batch of mallein PPD is tested for sterility, safety, preservatives, protein content and potency.
Sterility testing is performed according to the European Pharmacopoeia guidelines.
The examination for safety is conducted on from five to ten normal healthy horses by carrying out the intradermopalpebral test. The resulting swelling should be, at most, barely detectable and transient, without any signs of
conjunctival discharge.
Preparations containing phenol as a preservative should not contain more than 0.5% (w/v) phenol. The protein
content should be not less than 0.95 mg/ml and not more than 1.05 mg/ml.
Potency testing is performed in guinea-pigs and horses. The animals are sensitised by subcutaneous inoculation
with a concentrated suspension of heat-killed B. mallei in paraffin oil or incomplete Freund’s adjuvant. Cattle can
also be used instead of horses. The production batch is bioassayed against a standard mallein PPD by
intradermal injection in 0.1 ml doses in such a way that complete randomisation is obtained.
In guinea-pigs, the different areas of erythema are measured after 24 hours, and in horses the increase in skin
thickness is measured by calipers. The results are statistically evaluated, using standard statistical methods for
parallel-line assays.
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NIERMAN W.C., DESHAZER D., KIM H.S., TETTELIN H., NELSON K.E., FELDBLYUM T., ULRICH R.L., RONNING C.M.,
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S., SELENGUT J., SHAMBLIN C., SULLIVAN S.A., WHITE O., YU Y., ZAFAR N., ZHOU L. & FRASER C.M. (2004). Structural
flexibility in the Burkholderia mallei genome. Proc. Natl. Acad. Sci. USA., 101, 14246–14251.
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WERNERY U., KINNE J., ELSCHNER M. & NEUBAUER H. (2006). Glanders--a comprehensive review. Dtsch. Tierarztl.
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XIE X., XU F., XU B., DUAN X. & GONG R. (1980). A New Selective Medium for Isolation of Glanders Bacilli.
Collected papers of veterinary research. Control Institute of Veterinary Biologics, Ministry of Agriculture, Peking,
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YABUUCHI E., KOSAKO Y., OYAIZU H., YANO I., HOTTA H., HASHIMOTO Y., EZAKI T. & ARAKAWA M. (1992). Proposal of
Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new
genus with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. nov. Microbiol. Immunol.,
36, 1251–1275.
*
* *
NB: There are OIE Reference Laboratories for Glanders
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and diagnostic biologicals for glanders
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CHAPTER 2.5.12.
HORSE MANGE
See Chapter 2.9.8. Mange
*
* *
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.5.13.
VENEZUELAN EQUINE ENCEPHALOMYELITIS
SUMMARY
Venezuelan equine encephalomyelitis (VEE) viruses, of the genus Alphavirus of the family
Togaviridae, cause disease ranging from mild febrile reactions to fatal encephalitic zoonoses in
Equidae and humans. They are transmitted by haematophagous insects, primarily mammalophilic
mosquitoes.
The VEE complex of viruses includes six antigenic subtypes (I–VI). Within subtype I there are five
antigenic variants (variants AB–F). Originally, subtypes I-A and I-B were considered to be distinct
variants, but they are now considered to be identical (I-AB). Antigenic variants I-AB and I-C are
associated with epizootic activity in equids and human epidemics. Historically, severe outbreaks
have involved many thousands of human and equine cases. The other three variants of subtype I
(I-D, I-E, I-F) and the other five subtypes of VEE (II–VI) circulate in natural enzootic cycles. Equidae
serve as amplifying hosts for epizootic VEE strains while enzootic VEE viruses cycle primarily
between sylvatic rodents and mosquitoes. Enzootic variants and subtypes have been considered to
be nonpathogenic for equids, but can cause clinical disease in humans. During 1993 and 1996
however, limited outbreaks of encephalitis in horses in Mexico were shown to be caused by
enzootic VEE viruses of subtype I-E.
Identification of the agent: Diagnosis of VEE virus infection can be confirmed by the isolation,
identification, and antigenic classification of the isolated virus.
A presumptive diagnosis of equine encephalomyelitis can be made when susceptible animals in
tropical or subtropical areas display clinical signs of encephalomyelitis where haematophagous
insects are active. VEE virus can be isolated in cell cultures or in laboratory animals using the blood
or serum of febrile animals in an early stage of infection. It is recovered less frequently from the
blood or brain tissue of encephalitic animals.
VEE virus can be identified by complement fixation, haemagglutination inhibition, plaque reduction
neutralisation (PRN), or immunofluorescence tests using VEE-specific antibodies. Specific
identification of epizootic VEE variants can be made by the indirect fluorescent antibody test, or a
differential PRN test using subtype- or variant-specific monoclonal antibody, or by nucleic acid
sequencing.
Serological tests: Specific antibodies may be demonstrated by PRN tests against epizootic VEE
virus variants or by IgM capture enzyme-linked immunosorbent assay. Antibody can also be
demonstrated by the haemagglutination inhibition or the complement fixation tests.
Any diagnosis of VEE in an individual that is based on seroconversion in the absence of an
epizootic should be made with care. Infections of equids with enzootic VEE viruses produce a low
level viraemia accompanied by antibody development, but without clinical disease in most cases.
Antibody induced by such subclinical infections may be reactive to epizootic VEE virus variants.
Requirements for vaccines and diagnostic biologicals: The only acceptable vaccines against
VEE are an attenuated virus vaccine, made with strain TC-83, or inactivated virus preparations also
made from this strain. Attenuated virus is immunogenic when given by intramuscular injection, but
sometimes causes adverse reactions in the recipient.
Formalin-inactivated virulent VEE virus preparations should never be used in equids, as residual
virulent virus can remain after formalin treatment, and thereby cause severe illness in both animals
and humans. Epizootics of VEE have occurred from the use of such formalin-treated viruses.
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Chapter 2.5.13. — Venezuelan equine encephalomyelitis
A. INTRODUCTION
Venezuelan equine encephalomyelitis (VEE) is an arthropod-borne inflammatory viral infection of equines and
humans, resulting in mild to severe febrile and, occasionally fatal, encephalitic disease.
VEE viruses form a complex within the genus Alphavirus, family Togaviridae. The VEE virus complex is
composed of six subtypes (I–VI). Subtype I includes five antigenic variants (AB–F), of which variants 1-AB and 1C are associated with epizootic VEE in equids and concurrent epidemics in humans (Calisher et al., 1980; Monath
& Trent, 1981; Pan-American Health Organization, 1972; Walton, 1981; Walton et al., 1973; Walton & Grayson,
1989). The epizootic variants 1-AB and 1-C are thought to originate from mutations of the enzootic 1-D serotype
(Weaver et al., 2004); 1-AB and 1-C isolates have only been obtained during equine epizootics. The enzootic
strains include variants 1-D, 1-E and 1-F of subtype I, subtype II, four antigenic variants (A–D) of subtype III, and
subtypes IV–VI. Normally, enzootic VEE viruses do not produce clinical encephalomyelitis in the equine species
(Walton et al., 1973), but in 1993 and 1996 in Mexico, the 1-E enzootic subtype caused limited epizootics in
horses. The enzootic variants and subtypes can produce clinical disease in humans (Monath & Trent, 1981; PanAmerican Health Organization, 1972; Powers et al., 1997; Walton, 1981; Walton & Grayson, 1989).
Historically, epizootic VEE was limited to northern and western South America (Venezuela, Colombia, Ecuador,
Peru and Trinidad) (Pan-American Health Organization, 1972). From 1969 to 1972, however, epizootic activity
(variant 1-AB) occurred in Guatemala, El Salvador, Nicaragua, Honduras, Costa Rica, Belize, Mexico, and the
United States of America (USA) (Texas). Epizootics of VEE caused by I-AB or I-C virus have not occurred in
North America and Mexico since 1972. Recent equine and human isolations of epizootic VEE virus were subtype
1-C strains from Venezuela in 1993, 1995 and 1996 and Colombia in 1995.
The foci of enzootic variants and subtypes are found in areas classified as tropical wet forest, i.e. those areas with
a high water table or open swampy areas with meandering sunlit streams. These are the areas of the Americas
where rainfall is distributed throughout the year or areas permanently supplied with water. Enzootic viruses cycle
among rodents, and perhaps birds, by the feeding of mosquitoes (Monath & Trent, 1981; Pan-American Health
Organization, 1972; Walton, 1981; Walton & Grayson, 1989). Enzootic VEE strains have been identified in the
Florida Everglades (subtype II), Mexico (variant I-E), Central American countries (variant I-E), Panama (variants ID and I-E), Venezuela (variant I-D), Colombia (variant I-D), Peru (variants 1-D, III-C, and III-D), French Guiana
(variant III-B and subtype V), Ecuador (variant I-D), Suriname (variant III-A), Trinidad (variant III-A), Brazil
(variants I-F and III-A and subtype IV), and Argentina (subtype VI). In an atypical ecological niche, variant III-B
has been isolated in the USA (Colorado and South Dakota) in an unusual association with birds (Monath & Trent,
1981; Pan-American Health Organization, 1972; Walton, 1981; Walton & Grayson, 1989).
A tentative diagnosis of viral encephalomyelitis in equids can be based on the occurrence of acute neurological
disease during the summer in temperate climates or in the wet season in tropical or subtropical climates. These
are the seasons of haematophagous insect activity. Virus infection will result in clinical disease in many equids
concurrently rather than in isolated cases. Epizootic activity can move vast distances through susceptible
populations in a short time (Monath & Trent, 1981; Pan-American Health Organization, 1972; Walton, 1981;
Walton & Grayson, 1989). Differential diagnoses include eastern or western equine encephalomyelitis (chapter
2.5.5), Japanese encephalitis (chapter 2.1.7), West Nile fever (chapter 2.1.20), rabies (chapter 2.1.13) and other
infectious, parasitic, or non-infectious agents producing similar signs.
Human VEE virus infections have originated by aerosol transmission from the cage debris of infected laboratory
rodents and from laboratory accidents. Infections with both epizootic and enzootic variants and subtypes have
been acquired by laboratory workers (American Committee on Arthropod-Borne Viruses [ACAV], 1980). Severe
clinical disease or death can occur in humans. Those who handle infectious VEE viruses or their antigens
prepared from infected tissues or cell cultures should be vaccinated and shown to have demonstrable immunity in
the form of VEE virus-specific neutralising antibody (Berge et al., 1961; Pan-American Health Organization,
1972). All procedures producing aerosols from VEE virus materials should be conducted in biosafety cabinets at
containment level 3 (see Chapter 1.1.3 Biosafety and biosecurity in the veterinary microbiological laboratory and
animal facilities) (ACAV, 1980; United States Department of Health and Human Services, 1999).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
A confirmatory diagnosis of VEE is based on the isolation and identification of the virus or on the demonstration of
seroconversion. The period of viraemia coincides with the onset of pyrexia within 12–24 hours of infection.
Viraemia terminates 5–6 days after infection, and coincides with the production of neutralising antibodies and the
appearance of clinical neurological signs. Frequently, VEE viruses cannot be isolated from the brains of infected
equids. Blood samples for virus isolation should be collected from febrile animals that are closely associated with
clinical encephalitic cases.
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Chapter 2.5.13. — Venezuelan equine encephalomyelitis
Virus may be isolated from the blood or sera of infected animals by inoculating 1–4-day-old mice or hamsters
intracerebrally or by the inoculation of other laboratory animals, such as guinea-pigs and weaned mice. It may
also be isolated by the inoculation of various cell cultures including African green monkey kidney (Vero), rabbit
kidney (RK-13), baby hamster kidney (BHK-21), or duck or chicken embryo fibroblasts, or by inoculation of
embryonated chicken eggs. Details of virus identification techniques are described in chapter 2.5.5.
Isolates can be identified as VEE virus by complement fixation (CF), haemagglutination inhibition (HI), or plaque
reduction neutralisation (PRN) tests, or by immunofluorescence as described in chapter 2.5.5. The VEE virus
isolates can be characterised by the indirect fluorescent antibody or PRN tests using monoclonal antibody or by
nucleic acid sequencing. The VEE virus characterisation should be carried out in a reference laboratory (see
Table given in Part 4 of this Terrestrial Manual).
2.
Serological tests
Diagnosis of VEE virus infection in equids requires the demonstration of specific antibodies in paired serum
samples collected in the acute and convalescent phases. After infection, PRN antibodies appear within 5–7 days,
CF antibodies within 6–9 days, and HI antibodies within 6–7 days. The second convalescent phase serum sample
should be collected 4–7 days after the collection of the first acute phase sample or at the time of death. The
serological procedures are described in detail in chapter 2.5.5. Vaccination history must be taken into account
when interpreting any of the VEE serological test results. In horses not recently vaccinated with an attenuated live
virus strain, demonstration of VEE-specific serum IgM antibodies in a single serum sample supports recent virus
exposure.
Any diagnosis of VEE in an individual that is based on seroconversion in the absence of an epizootic should be
made with care. Although enzootic subtypes and variants are nonpathogenic for equids, infection will stimulate
antibody production to epizootic VEE virus variants.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
The acceptable vaccines against VEE infection are an attenuated virus vaccine, strain TC-83, and an inactivated
virus preparation made from that strain (Monath & Trent, 1981; Pan-American Health Organization, 1972; Walton,
1981; Walton & Grayson, 1989). The inactivated vaccine is now the most widely used, and is marketed in
EEE/VEE, EEE/WEE/VEE, EEE/WEE//VEE/tetanus toxoid, and EEE/WEE/VEE/West Nile virus/tetanus toxoid
combinations.
Inactivated vaccine should be administered in two doses with an interval of 2–4 weeks between doses. Annual
revaccination is recommended.
Attenuated vaccine should be reconstituted with physiological saline and used immediately. Multidose vials are
kept on ice while the vaccine is being used. Any vaccine not used within 4 hours of reconstitution should be safely
discarded. Foals under 2 weeks of age and pregnant mares should not be vaccinated. Animals are vaccinated
subcutaneously in the cervical region with a single dose. Revaccination is not recommended.
NOTE: Formalin-treated preparations of virulent epizootic VEE virus should never be used in equids. Residual
virulent virus can remain after formalin treatment, and result in severe illness. Epizootics of VEE have occurred in
Central and Southern America from the use of such preparations (Walton, 1981; Weaver et al., 1999).
1.
Seed management1
a)
Characteristics of the seed
Attenuated VEE virus vaccine strain TC-83 originated from the Trinidad donkey strain (a variant of I-AB) of
epizootic VEE virus isolated in 1944. This strain was derived by serial passage of the Trinidad donkey strain
in fetal guinea pig heart cells. It is safe and immunogenic at the established passage levels, and induces
protective immunity in vaccinated equids, although adverse reactions can sometimes occur. The vaccine
was originally developed for use in personnel involved in high-risk VEE virus research. Suitable seed lots
should be maintained at –70°C in a lyophilised state.
1
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Sections on Seed management, Manufacture, In-process control, and Batch control are taken from the Biotechnology,
Biologics, and Environmental Protection Division of the United States Department of Agriculture’s Animal and Plant Health
Inspection Service (APHIS).
OIE Terrestrial Manual 2012
Chapter 2.5.13. — Venezuelan equine encephalomyelitis
b)
Method of culture
The virus is grown in fetal guinea pig heart cell cultures in a suitable medium.
c)
Validation as a vaccine
The cells used for vaccine production must be free from bacterial, fungal, mycoplasmal, and viral
contamination. VEE virus is identified in batches of vaccine by PRN tests against hyperimmune serum. For
inactivated vaccines of cell culture origin, strain TC-83 virus is treated with formaldehyde.
2.
Method of manufacture (see footnote 1)
Vaccine is produced by harvesting supernatant fluid from fetal guinea pig heart monolayers in which the
replication of attenuated VEE virus has occurred. The monolayers are maintained at approximately 37°C. The
time of harvesting is determined by the occurrence of characteristic cytopathic changes when approximately 70–
100% of the cell sheet is affected, typically 1–3 days after infection. The supernatant fluid is clarified by low speed
centrifugation and suitable stabilisers are added to protect the virus during freezing and lyophilisation.
3.
In-process control (see footnote 1)
Cultures should be examined daily for cytopathic changes. After harvesting, the virus suspension should be
tested for the presence of microbial contaminants.
Inactivated vaccines derived from attenuated strain TC-83 virus should be checked to exclude the presence of
viable virus after formalin treatment.
4.
Batch control (see footnote 1)
a)
Sterility
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
b)
Safety
Safety tests of the inactivated vaccine are described in chapter 2.5.5.
Safety tests of the attenuated vaccine are conducted in mice. A 0.5 ml dose is injected intraperitoneally or
subcutaneously into each of eight mice, and the animals are kept under observation for 7 days. If adverse
reactions attributable to the product occur during this period, the product is considered to be unsatisfactory.
c)
Potency
Potency tests of the inactivated vaccine are described in chapter 2.5.5 except that antibody titre in
inoculated guinea-pigs will be ≥1/4.
Potency of the attenuated vaccine can be determined by testing in horses. Each of 20 susceptible horses is
inoculated subcutaneously with 1 ml of lyophilised vaccine that has a reconstituted virus titre of at least
2.5 log10 TCID50 (50% tissue culture infective dose) per ml. For a valid test, at least 19 of 20 vaccinated
horses must have HI antibody titres of at least 1/20 or serum neutralising antibody titres of at least 1/40
within 21–28 days of vaccination.
When tested at any time within the expiration period following lyophilisation, the product must have a virus
titre of 0.7 log10 greater than that used to test horses as described above, but no less than
2.5 log10 TCID50/dose.
The final product must be free from bacterial, fungal, mycoplasmal, or extraneous viral contaminants.
d)
Duration of immunity
Comprehensive studies on duration of immunity are not available. An annual revaccination is recommended
for the inactivated vaccine. Foals that are vaccinated at under 1 year of age should be revaccinated before
the next vector season. Revaccination with the attenuated vaccine is not recommended.
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Chapter 2.5.13. — Venezuelan equine encephalomyelitis
e)
Stability
The lyophilised vaccine is stable and immunogenic for 3 years if kept refrigerated at 2–7°C. After 3 years,
vaccine should be discarded. The vaccines should be used immediately after reconstitution. Multidose vials
of the attenuated vaccine should be kept on ice while being used. All unused vaccine should be safely
discarded 4 hours after reconstitution.
f)
Preservatives
The preservatives used are thimerosal at a 1/1000 dilution and antibiotics (neomycin, polymyxin,
amphotercin B, and gentamicin).
g)
Precautions (hazards)
Pregnant mares and foals under 2 weeks of age should not be vaccinated.
All personnel handling infectious VEE viruses or their antigens prepared from infected tissues or cell cultures
should be vaccinated and shown to have demonstrable immunity in the form of VEE virus-specific
neutralising antibody. All procedures producing aerosols from VEE virus materials should be conducted in
biosafety cabinets with biocontainment and efficient filtration of the exhaust air from the laboratory (ACAV,
1980; United States Department of Health and Human Services, 1999).
5.
Tests on the final product
a)
Safety and potency
Safety and potency tests are as outlined above under Batch control (Sections C.4.b and C.4.c). The
attenuated vaccine must have a virus titre of no less than 2.5 log10 TCID50/dose.
REFERENCES
AMERICAN COMMITTEE ON ARTHROPOD-BORNE VIRUSES (ACAV), SUBCOMMITTEE ON ARBOVIRUS LABORATORY SAFETY
(1980). Laboratory safety for arboviruses and certain viruses of vertebrates. Am. J. Trop. Med. Hyg., 29, 1359–
1381.
BERGE T.O., BANKS I.S. & TIGERTT W.D. (1961). Attenuation of Venezuelan equine encephalomyelitis virus by in
vitro cultivation in guinea pig heart cells. Am. J. Hyg., 73, 209–218.
CALISHER C.H., SHOPE R.E., BRANDT W., CASALS J., KARABATSOS N., MURPHY F.A., TESH R.B. & WIEBE M.E. (1980).
Proposed antigenic classification of registered arboviruses. I. Togavirus Alphavirus. Intervirology, 14, 229–232.
MONATH T.P. & TRENT D.W. (1981). Togaviral diseases of domestic animals. Comp. Diagn. Vir. Dis., 4, 331–440.
PAN-AMERICAN HEALTH ORGANIZATION (1972). Venezuelan encephalitis. In: Proceedings of a Workshop/
Symposium on Venezuelan Encephalitis Virus. Sci. Publ. No. 243, Washington DC, USA, 416 pp.
POWERS A.M., OBERSTE M.S., BRAULT A.C., RICO-HESSE R., SCHMURA S.M., SMITH J.F., KANG W, SWEENEY W.P. &
WEAVER S.C. (1997). Repeated emergence of epidemic/epizootic Venezuelan equine encephalitis from a single
genotype of enzootic subtype ID virus. J. Virol., 71, 6697–6705.
UNITED STATES DEPARTMENT OF HEALTH AND HUMAN SERVICES (1999). Biosafety in Microbiological and Biomedical
Laboratories. US Government Printing Office, Washington DC, USA, 190–196.
WALTON T.E. (1981). Chapter 24. Equine encephalomyelitis. In: Virus Diseases of Food Animals. A World
Geography of Epidemiology and Control. Disease Monographs, Vol. 2, Gibbs E.P.J., ed. Academic Press, New
York, USA, 587–625.
WALTON T.E., ALVAREZ O. Jr, BUCKWALTER R.M. & JOHNSON K.M. (1973). Experimental infection of horses with
enzootic and epizootic strains of Venezuelan equine encephalomyelitis virus. J. Infect. Dis., 128, 271–282.
WALTON T.E. & GRAYSON M.A. (1989). Chapter 46. Venezuelan equine encephalomyelitis. In: The Arboviruses:
Epidemiology and Ecology, Vol. 4, Monath T.P., ed. CRC Press, Boca Raton, Florida, USA, 203–231.
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WEAVER S.C., FERRO C., BARREREA R. BOSCHELL J. & NAVARRO J.C. (2004) Venezuelan equine encephalitis. Annu.
Rev. Entomol., 49, 141–174.
WEAVER S.C., PFEFFER M., MARRIOTT K., KANG W. & KINNEY R.M. (1999) Genetic evidence for the origins of
Venezuelan equine encephalitis virus subtype IAB outbreaks. Am. J. Trop. Med. Hyg., 60, 441–448.
*
* *
NB: There is an OIE Reference Laboratory for Venezuelan equine encephalomyelitis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for Venezuelan equine encephalomyelitis
OIE Terrestrial Manual 2012
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
SECTION 2.6.
LEPORIDAE
CHAPTER 2.6.1.
MYXOMATOSIS
SUMMARY
Myxomatosis is a major disease of the European rabbit caused by the Myxoma virus, a member of
the Poxviridae family. The diagnosis of myxomatosis, regardless of its clinical form, depends on the
isolation and identification of the virus or the demonstration of its antigens. The presence of a
humoral immune response facilitates a retrospective diagnosis of a mild form of the disease, and
can provide an indication of the prevalence of infection in a rabbit population. The disease is
characterised by gross myxomatous skin lesions.
Identification of the agent: When skin lesions are present on a dead rabbit, the viral antigen may
be demonstrated by immunodiffusion tests on lesion fragments. Monolayer cell cultures of rabbit
kidney inoculated with lesion material will show the characteristic cytopathic effects of poxviruses.
The presence of virus can be confirmed by immunofluorescence and negative-staining electron
microscopy.
The inoculation of rabbits with suspect material takes longer to identify infection, but this will serve
to confirm the presence of infective virus and indicate its pathogenicity.
Serological tests: Identification and titration of specific antibodies arising from natural infection or
from immunisation are done by traditional complement fixation or by a recently developed and more
sensitive enzyme-linked immunosorbent assay (ELISA), which is not affected by pro- or anticomplementary factors. The difficulty in obtaining blood samples from representative members of a
population can be obviated by collecting blood dried on filter paper; this can later be extracted and
examined by the indirect fluorescent antibody test or ELISA. Microcapillary blood sampling can also
be used for the ELISA.
Qualitative agar gel immunodiffusion tests have the advantage of detecting both antigen and
humoral antibodies.
Requirements for vaccines and diagnostic biologicals: Modified live virus vaccines prepared
from fibroma virus or modified Myxoma virus strains are available for immunisation of rabbits.
A. INTRODUCTION
Myxomatosis is a major viral disease of wild and domestic European rabbits (Oryctolagus cuniculus) caused by
the Myxoma virus (MV), a member of the Poxviridae family. The aetiological agent was first isolated from a colony
of laboratory rabbits in Uruguay in 1898 and identified as a poxvirus in 1927. The natural hosts are two species of
leporid: Sylvilagus brasiliensis in South America (South American strains) and Sylvilagus bachmani (Californian
strains) in California (Fenner, 1994). In its natural hosts, the viral strains produce only a benign fibroma,
generalised disease occurring only in juvenile animals. In the European rabbits, two forms of the disease have
been identified to date: the nodular (classical) form and the amyxomatous (respiratory) form.
Florid skin lesions and severe immunodysfunction, accompanied by supervening Gram-negative bacterial
infections of the respiratory tract, characterise the nodular myxomatosis syndrome caused by a virulent MV strain.
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Chapter 2.6.1. — Myxomatosis
Prototype strains of virus deriving from the Australian and European outbreaks have been designed that
characterise the various virulence grades (from grade I to grade V) as determined in laboratory rabbits (Fenner &
Ratcliffe, 1965). After infection with a grade I (the most virulent) strain, the first sign of infection is a lump at the
site of infection, which increases in size and usually becomes protuberant and ulcerates. An acute blepharoconjunctivitis and an oedematous swelling of the perineum and scrotum gradually develop. The secondary skin
lesions appear on about the sixth or the seventh day (Fenner, 1994). Death usually occurs between the eighth
and fifteenth day post-infection. After infection with grade II to V strains, the clinical signs are usually the same
with the exception that they evolve more slowly and are less severe. When animals survive, the lesions
progressively heal. The mortality rate fluctuates between 20 and 100%, according to the viral strain. The natural
mode of transmission of the nodular form is by biting insects, but limited transmission from rabbit to rabbit is
possible if they are closely confined. This form is mainly observed in small-scale rabbitries (Arthur & Louzis,
1988). The clinical signs of amyxomatous myxomatosis are mainly respiratory, skin nodules being few and small.
It might be thought that amyxomatous myxomatosis would not spread via vectors but through direct contact, and
would arise predominantly in intensive enclosed rabbitries. However, this last notion must be viewed with caution
as the disease has also been observed in wild rabbits. So far, these forms of myxomatosis have been reported
only in France (Brun et al., 1981; Joubert et al., 1982), Spain (Rosell et al., 1984) and more recently in Belgium
(Marlier & Vindevogel, 1996).
As MV has a very narrow host range (it only infects leporides), there is no health risk to humans.
B. DIAGNOSTIC TECHNIQUES
As the signs of the disease become less distinct with the attenuation of virus strains, the submission of samples
for laboratory diagnosis becomes more important. Moreover, the expression of the ectodermotropism is clearly
reduced for amyxomatous MV strains, and the clinical diagnosis of the amyxomatous form of myxomatosis is
clearly more difficult than for the classical one. The different techniques available vary in their ability to detect MV
in typical myxomatous lesions, oedema of the eyelids or genital oedema. Nevertheless, the diagnosis of
attenuated typical myxomatosis or of atypical (amyxomatous) forms most often requires the isolation of the virus
by inoculation of sensitive cell lines such as the RK-13 cell line (ATCC CCL37) and identification of the virus by
immunological methods. In both cases, the agent can also be identified by demonstration of MV nucleic acid by
polymerase chain reaction, molecular techniques were not specifically evaluated for diagnosis.
1.
Identification of the agent
A portion of lesion (myxoma or pieces of organs or tissues, especially eyelids) is excised with scissors. Myxoma
are separated from the epidermis and superficial dermis. This is washed with phosphate buffered saline (PBS)
with antibiotics as defined below and homogenised with ground glass at a dilution rate of 1 g tissue/4.5–9.0 ml of
PBS. Cells are disrupted by two freeze–thaw cycles, or by ultrasonication to liberate virions and viral antigens.
This suspension is centrifuged for 5–10 minutes at 1500 g. The supernatant fluid is used for the tests.
a)
Culture
Isolation of the virus in cell culture is accomplished using primary cultures of rabbit kidney (RK) cells, or with
established cell lines, such as RK-13, in Opti-MEM containing 2% calf serum, 300 international units (IU)/ml
penicillin; 300 µg/ml streptomycin; 100 µg/ml gentamycin; 50 IU/ml nystatin (mycostatin); and 5 µg/ml
amphotericin (fungizone). The inoculum consists of the supernatant fluid from a homogenised lesion or
oculo-respiratory discharge in Opti-MEM with 2% calf serum and antibiotics. This is removed from the cell
layer after 2 hours. The cell layer is washed in a small volume of medium and then replenished with
maintenance medium (Opti-MEM).
A cytopathic effect (CPE) typical of poxviruses (Joubert, 1973) usually develops after 24–48 hours (37°C
and 5% CO2), but with some strains, it may take up to 7 days for CPE to be observed. According to the viral
strain, groups of cells with a confluent cytoplasm form syncytia that vary in size from 2 to 50 or even
100 nuclei together. The nuclei of some cells change, the chromatin forming basophilic aggregations that
vary in size and number and give the culture a leopard-skin appearance. Eosinophilic intracytoplasmic
inclusions remain discrete, if present at all. Affected cells round up, contract and become pyknotic. They
then lyse and become detached from the glass or plastic support. Later, all cells are affected and the cell
monolayer detaches completely.
Shope’s fibroma virus at first produces well-defined voluminous masses of rounded cells, which proliferate
and pile up (Joubert, 1973). At the edge, cells just becoming infected show discrete nuclear changes and
acidophilic cytoplasmic inclusions that are numerous at an early stage. The cell layer is destroyed after
several days.
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Chapter 2.6.1. — Myxomatosis
b)
Immunological methods
Agar gel immunodiffusion (AGID) tests are simple and rapid to perform – results can be obtained within
24 hours. Agar plates are prepared with Noble agar (0.6 g), ethylene diamine tetra-acetic acid (EDTA)
(2.5 g), sodium chloride (4.5 g), and distilled water (500 ml) containing thiomersal (merthiolate) at 1/100,000
dilution. Standard antiserum (see below), and the test sample are placed in opposing wells that are 6 mm in
diameter and 5 mm apart. Another technique is to deposit a small portion of the lesion directly into the agar,
5 mm away from a filter paper disk impregnated with the antiserum. A number of lines of precipitation,
usually up to three, appear within 48 hours, indicating the presence of myxoma viral antigens. Only one line
forms in the presence of heterologous reactions with Shope’s fibroma virus.
Indirect fluorescent antibody (IFA) tests can be applied to cultures from 24 hours onwards. IFA tests reveal
intracytoplasmic multiplication of virus, without being able to distinguish MV from Shope’s fibroma virus. The
inoculation of chicken embryo cells (trypsinised at day 11 of egg incubation) does not result in CPE, but it is
useful for detecting the viral antigens by IFA tests.
c)
Electron microscopy
Negative-staining electron microscopy (EM) can be applied to a portion of skin lesion. The technique is
simple and rapid to perform, giving results in 1 hour. About 1 mm3 of the tissue is laid in a watch glass and
three drops of distilled water are added. After 1–2 minutes at room temperature an EM grid coated with
formvar and carbon is laid over the liquid. After 1 minute any excess liquid is removed with filter paper and
immediately a 2% aqueous solution of ammonium molybdate, pH 7.0, is dropped on to the grid. After
10 seconds the excess liquid is removed with filter paper and the grid is prepared for the electron
microscope. In a positive case, typical poxvirus particles can be seen. MV cannot be distinguished from
Shope’s fibroma virus using this method.
d)
Inoculation tests
Rabbit intradermal inoculation also offers a means of identifying the virus through its special characteristics
and pathogenicity (virulence grade, classical or amyxomatous forms). It should be avoided if possible but
has the advantage of being a gauge of virulence, from the type of inflammation in lesions (local or systemic
infection) to the extent of lesions and survival time, and can distinguish Shope’s fibroma virus (with its simple
fibromatous local lesion) from MV (capable of causing generalised infection in adults). Rabbits should be of
a domestic breed, weighing approximately 2 kg, unvaccinated and previously tested for the absence of
antibodies (Joubert, 1973).
The inoculum may be the supernatant fluid from a homogenised lesion (with antibiotics) or the product of a
cell culture. Between 0.1 and 0.2 ml is administered intradermally behind the ear or into the dorso-lumbar
region, which has previously been depilated. The inoculum may be assayed by injecting serial dilutions in
saline buffer at one site for each dilution. A primary lesion will appear at the sites within 2–5 days, followed
by conjunctivitis. Using five sites for each dilution allows a 50% infective dose (ID50) to be obtained. If the
animal survives, the disease can be confirmed serologically after 15 days.
2.
Serological tests
Antibodies develop within 8–13 days. In the nonlethal forms and in vaccinated rabbits, the titre is highest after 20–
60 days; it declines thereafter, disappearing after 6–8 months in the absence of reinfection (serological response
evaluated by use of the complement fixation [CF] test) (Saurat et al., 1980).
Various serological tests may be used, but agar gel immunodiffusion (AGID), CF, IFA and enzyme-linked
immunosorbent assay (ELISA) (in order of increasing sensitivity) are the most appropriate tests for international
trade and other applications. These tests require standard antigens and antisera. The antigen can be prepared
from the Lausanne strain, or some antigenically related strain, propagated in rabbits or cell cultures.
•
Preparation of standard reagents (AGID, CF and IFA tests)
•
Preparation of antigen
Myxomatous lesions are removed from rabbits at 6–7 days after inoculation and homogenised in veronal
buffer to a dilution of 1/5. The antigen is the supernatant fluid that is obtained following centrifugation (5–
10 minutes, 1500 g). Any anticomplementary activity is abolished by adding 0.6% chloroform. The antigen
fluid can be frozen at–30°C or –70°C for stock purposes or used directly in CF tests after titration against a
standard antiserum.
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Antigen is made from cell cultures using the RK-13 cell line. RK-13 (rabbit kidney cells CCL-37) are cultured
in Dulbecco’s modified Eagle’s medium (DMEM, Gibco) containing penicillin and streptomycin and 10% fetal
calf serum (FCS) for 48 hours before infection. Cells are cultured at 37°C in 5% CO2. Cells are infected by
the MV strain at a multiplicity of infection (m.o.i.) of 1–5. After incubation for 2 hours at 37°C, the inoculum is
removed and the cells are maintained in DMEM with 5% FCS for 48 hours at 37°C with 5% CO2. The
monolayer is harvested about 48 hours after infection, when the cells clearly show CPE (80%), and is
centrifuged (1000 g). The supernatant fluid is retained. Infected cells are frozen and thawed three times to
release additional virus and the viral suspension is clarified at 1000 g. The newly harvested supernatant is
added to the original supernatant. The final supernatant fluid is the antigen, and is stored at –20°C or –70°C
(for longer conservation). It is titrated in cell cultures before use.
•
Titration of standard antigen by complement fixation test
i)
Inactivate standard antiserum for 30 minutes in a water bath at 56°C. No international standard serum
for myxomatosis is available, but internal positive standards should be prepared and the titre estimated
in the appropriate range, using the CF test or ELISA. After this, the following procedure is used to
standardise batches of antigen.
ii)
Make doubling dilutions of standard serum in calcium/magnesium/veronal buffer (CMV)
(BIOMERIEUX, ref 72171), pH 7.2, from 1/2 to 1/4096, using a 96-well round-bottomed microtitre plate,
one row (column) per dilution and 25 µl per well.
iii)
Using tubes, make doubling dilutions of antigen in CMV, from 1/10 to 1/1280.
iv)
Add 25 µl of the first antigen dilution to each well in the first line of the plate. Similarly add succeeding
dilutions of antigen to subsequent lines of the plate so that a checkerboard titration of antigen and
antibody is created.
v)
Add 25 µl (6 H50 units [50% haemolysis]) of complement per well.
vi)
Incubate the plate, covered with a plastic film, for 1 hour at 37°C or 14 hours at 4°C.
vii)
Add 50 µl per well of the haemolytic system (2.5% sheep red blood cells [RBCs] and an equal volume
of anti-sheep RBC serum diluted both in CMV) (for optimal working dilution of anti-sheep RBC, follow
the recommendations of the producer; alternatively it should be determined individually for each lot of
serum used).
viii) Cover the plate again and incubate for 30 minutes at 37°C.
ix)
•
•
Read the highest dilution of antigen giving complete haemolysis (H100) with the highest dilution of
standard serum. There is 1 antigenic unit (AgU) in 25 µl of antigen of this dilution.
Myxoma virus titration of infectious particles in cell cultures
i)
MV is diluted from 10–1 to 10–5 in DMEM + antibiotics + 2% FCS.
ii)
Confluent RK-13 cells in P6 (Falcon plates) are infected in triplicate with 200 µl of serial dilutions of
interest and incubated at 37°C for 2 hours in a 5% CO2 incubator.
iii)
Inoculum is then removed and 2 ml of DMEM + 5% FCS is added to each well.
iv)
Incubation is performed at 37°C in a 5% CO2 incubator for 2 days.
v)
Medium is then removed and replaced by solid medium with 1% LMP agarose and 2% FCS.
vi)
Incubation is performed for 1 or 2 days more, then plaques are counted without coloration for each
dilution (coloration is possible with an overlay containing neutral red).
vii)
After average calculation, the titre of the viral suspension is expressed in plaque-forming units (PFU)
per ml.
Preparation and titration of standard positive serum
For the standard antiserum, an adult serologically negative rabbit is vaccinated with an attenuated strain of
MV, or with the Shope’s fibroma virus. After 3–4 weeks, the rabbit is inoculated intradermally with virulent
myxoma virus (Lausanne strain or a related one) (5 × 103 PFU). Serum is obtained 3 weeks later and
titrated by the CF test or ELISA. If the titre is >1/640 or > 1/1000, respectively for CF test and ELISA, the
animal is bled and the serum is stored at –20°C.
a)
Complement fixation test
CF tests (Saurat et al., 1980) are done in tubes or in microtitre plates (Chantal et al., 1993) by conventional
methods, recording 100% or 50% haemolysis. This is the standard method at the present time.
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i)
Titrate the complement in haemolysis tubes, in the presence of 1 AgU, in order to determine the
H50 unit.
ii)
Inactivate the test serum and the positive and negative control sera in a water bath for 30 minutes at
56°C.
iii)
Make doubling dilutions of test and control sera in CMV, from 1/4 to 1/1024, using a 96-well roundbottomed microtitre plate and 25 µl per well. Use the first well for the initial 1/4 dilution and the second
as a serum control (anticomplementary control at 1/4 dilution). Provide antigen (without serum),
complement, and RBC control wells (see below) (two wells of each).
iv)
Add 1 AgU of MV antigen in 25 µl per well (except to serum, complement and RBC control wells), then
add 6 H50 of complement in 25 µl per well (except to RBC control wells).
v)
Incubate the plates, covered with plastic film, for 14 hours (overnight) at 4°C.
vi)
Add 50 µl per well of the haemolytic system
vii)
Cover the plates again and incubate for 30 minutes at 37°C.
viii) Prepare H100, H75, H50, H25 haemolysis controls using complement controls (H100) and CMV.
b)
ix)
Read after centrifugation (1000 g, 10 minutes) or passive sedimentation at 4°C. The test sera results
are determined as the highest dilution of serum that gives at least 50% haemolysis inhibition.
x)
A negative serum should give haemolysis inhibition <50% at 1/4 dilution.
Indirect fluorescent antibody test
IFA test (Gilbert et al., 1989) is carried out using chicken embryo or RK-13 cell cultures in flat-bottomed wells
of microtitre plates: cell s suspension, 4 × 104 cells diluted in medium, is distributed into all wells and a
confluent cell sheet is formed within 24 hours. The medium is discarded and 100 µl of viral suspension (with
a multiplicity of infection of 0.05) is added to each well. After 2 hours, 100 µl of Minimal essential medium
(MEM) containing 2% calf serum is added. After 48 hours of incubation, the plates are washed with PBS and
fixed with acetone containing 50% ethanol for 30 minutes at –20°C, or paraformaldehyde (4% in PBS) at
room temperature. The plates are then dried at 37°C for 15 minutes. The plates can be stored at –30°C or
–70°C for 3 months. Sera are tested by IFA using anti-rabbit IgG conjugated to fluorescein isothiocyanate.
The test results may be qualitative with sera diluted 1/20, or quantitative with serial dilutions of serum.
c)
Enzyme-linked immunosorbent assay
An ELISA has been developed (Gelfi et al., 1999) that uses a semi-purified myxomatosis virus (French
hypervirulent T1 strain antigenically related to Lausanne strain) produced in RK-13. The virus is harvested
as a suspension of cells 48 hours after infection, and is centrifuged. The cell pellet is homogenised in TL20
(20 mM Tris, pH 8.6, 150 mM NaCl, and 1 mM EDTA), disrupted in ground glass and centrifuged at 1200 g
at 4°C for 10 minutes.
The supernatant fluid is laid down on an equal volume of a 36% sucrose cushion in TL20 and centrifuged at
200,000 g for 2 hours in an SW 41 rotor at 4°C. The pellet is homogenised in 4–12 ml TL20 and again laid
down on a 36% sucrose cushion.
The new pellet is homogenised in 0.5–1 ml TL20 and quantified by the Bradford method (colorimetric
reaction with Coomassie brilliant blue) (Bertagnoli et al., 1996) or spectrophotometry (viral proteins account
for around three-fifths of total protein). It can be stored at –30°C before use.
i)
Coat probind (Falcon) assay plates for 16 hours (overnight at 37°C) with 1 µg per well viral proteins in
100 µl PBS, pH 7.6, leaving one column blank (i.e. coat with PBS only). Note that different batches of
antigen vary in activity, and should be titrated against known standards to select antigen with high
optical densities (OD).
ii)
After three washes in PBS, block free binding sites by incubation in 25 mg/ml gelatin in PBS for 1 hour
at 37°C.
iii)
Wash the plate three times in PBS–0.01% Tween 20, and add 100 µl serial twofold dilutions of serum
in PBS–Tween. Include positive and negative serum standards in each plate.
iv)
After 60 minutes’ incubation at 37°C and three washes in PBS–Tween, add 100 µl of a dilution in PBS–
Tween of goat anti-rabbit IgG serum (previously tested) conjugated to alkaline phosphatase for 1 hour
at 37°C.
v)
After four washes in PBS–Tween and one more in PBS alone add, as substrate, 100 µl of disodium pnitrophenyl phosphate at a concentration of 1 mg/ml in 10% diethanolamine.
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vi)
After 12 minutes in the dark at room temperature, the enzymatic reaction is stopped by the addition of
50 µl of 2 N NaOH.
vii)
Read OD in a spectrophotometer at a wavelength of 405 nm.
viii) Express the serum sample titre as the inverse of the highest dilution for which the OD value is bigger
than three times the OD value of the negative serum standard at the same dilution.
The detection of specific MV antibodies by ELISA has been shown to be a highly sensitive and specific
method for kinetic studies in experimental infection (Bertagnoli et al., 1996). Evaluation of the test has shown
its great value for diagnostic application in wild rabbit populations (Gelfi et al., 1999).
For epidemiological surveys, the IFA test and the indirect ELISA can also be carried out using blood dried on
blotting or filter paper: discs are cut (paper punch size) and two discs are placed in each well, to which is
added 100 µl PBS to extract the serum. The extract product is about 1/20 diluted and can be used as a fresh
sample for testing (Gilbert et al., 1989). Blood samples collected in anticoagulant-coated capillary tubes can
be used for the ELISA. The sample is washed in the diluting solution to obtain the required dilution (Rodak et
al., 1985).
d)
Agar gel immunodiffusion test
Agar gel immunodiffusion (AGID) (Sobey et al., 1966) is qualitative and can detect antigen or antibody. Agar
is prepared as described previously (Section B.1) using 6 ml per 10 cm Petri dish. Strips of filter paper
containing the standard antigen and antiserum, and discs containing test sera are arranged on the surface of
the agar (discs between the strips). The plates are incubated in a humid atmosphere at 37°C and read after
24–48 hours. Three precipitation lines should appear. If the test sera contain MV-specific antibody, at least
one of the three lines is bent towards the antigen band; otherwise it remains straight. If sera contain MV
antigen, at least one of the lines is bent towards the standard serum strip. The test can also be carried out in
a more conventional manner using liquid reagents in wells cut in the agar.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
Two types of live vaccine are used for vaccination against myxomatosis: a heterologous vaccine prepared from
Shope’s fibroma virus (Fenner & Woodroofe, 1954; Jobert, 1983; Shope, 1932), and a homologous vaccine
prepared from an attenuated strain of MV (Arguello Villares, 1986; Gorski & Mizak, 1985; Saurat et al., 1978;
Tozzini & Mani, 1975; Von Der Ahe et al., 1981). They are administered subcutaneously or intradermally.
A new recombinant MV expressing rabbit haemorrhagic disease virus (RHDV) capsid protein and conferring
double protection against myxomatosis and RHDV (Bertagnoli et al., 1996) has been developed, but is not yet
available commercially.
Guidelines for the production of veterinary vaccines are given in Chapter 1.1.6 Principles of veterinary vaccine
production. The guidelines given here and in chapter 1.1.6 are intended to be general in nature and may be
supplemented by national and regional requirements.
A master seed virus (MSV) must be established and used according to a seed-lot system. A record must be kept
of its origin, passage history and characteristics.
1.
Seed management
a)
Characteristics of the seed
The viruses employed are Shope’s fibroma virus or MV. The strains of Shope’s fibroma virus are usually the
original Shope’s OA strain (1932), Boerlage’s strain or various closely related strains. The strains of MV are
modified by passaging in embryonated chicken eggs, rabbit kidney cells at decreasing temperatures, or
chicken embryo cells. The strains usually result from having been cloned several times.
b)
Method of culture
Shope’s fibroma virus strains are maintained by passage in specific pathogen free (SPF) rabbits or in
unvaccinated rabbits from a stock known to be free from myxomatosis. Skin on the backs of healthy adult
rabbits is shaved, and multiple sites are inoculated with a 1% suspension of virulent material. Fibromas are
fully developed within 8–10 days, at which time the rabbits are killed and the tumours are removed
aseptically and homogenised with distilled water. The suspension is stored at –30°C or –70°C in 50%
buffered glycerol, or as a 5% dilution in a protein solution (bovine albumin). The production of the Shope’s
fibroma virus is also possible in rabbit dermal cell line.
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MV is grown on chicken embryo cell culture obtained from flocks free from specified pathogens, or on
suitable cell lines (rabbit dermal cell line). Virus can also be cultivated on RK-13 cells.
c)
Validation as a vaccine
i)
Identity
Specific antigenic characteristics of the Shope’s fibroma virus strains are verified by AGID using sera
against fibroma and myxomatosis.
The identity of MV is confirmed by neutralisation tests in RK-13 cells, or in a suitable cell line using a
monospecific antiserum (produced by vaccination of rabbits with the specific vaccine viral strain).
ii)
Purity
The Master seed must be free from bacterial, fungal, mycoplasmal and viral contamination.
iii)
Safety
Samples for safety testing are taken from a batch produced according to the manufacturing process.
The dose to be used shall contain the maximum titre or potency established by the manufacturer
(release titre).
Several tests are performed, at the Master Seed level, to demonstrate different aspects of safety. The
safety of 10 times the normal dose must be demonstrated. Also, it is necessary to examine the
dissemination of vaccine virus within the vaccinated animal, the ability of vaccine virus to spread from
the vaccinated animal to in-contact animals and to test whether there is reversion-to-virulence of the
vaccine virus, following serial passage in rabbits.
The pathogenicity of the Shope’s fibroma virus strains is tested by inoculating rabbits with serial
dilutions of supernatant fluids obtained by centrifugation of tumour preparations. Macroscopic and
histopathological features and the course of development of fibromas are tested in SPF rabbits
periodically. (Numerous serial passages in rabbits may induce mutation to the inflammatory IA strain,
which produces severe lesions that are more inflammatory than neoplastic.)
The residual pathogenicity of the MV strains is tested by intradermal inoculation into SPF rabbits or
unvaccinated rabbits free from myxomatosis. These rabbits should not develop more than a local
reaction with perhaps small secondary lesions on the head that disappear within a few days.
For both strains, the rectal temperature and the body weight should be recorded as additional
parameters.
iv)
Efficacy
Different trials must be undertaken from representative batches of final product containing the minimum
titre or potency. The protective effect is demonstrated as follows:
A minimum of ten adult rabbits are inoculated with a dose of fibroma vaccine, and three rabbits serve
as unvaccinated controls. After 14 days, all rabbits are inoculated, intradermally into the eyelids, with a
pathogenic strain of MV (example: 0.1 ml inoculum containing 103 ID50 [median infectious dose]).
During the following 21 days, the controls will die from myxomatosis, and at least seven of the ten
vaccinated rabbits must present no signs of generalised infection.
Similarly, myxoma vaccine is tested in ten rabbits with three controls. After 14 days, all the rabbits are
challenged with a sufficient quantity of virulent strain (example: 0.1 ml of the Lausanne virus strain
containing 103 ID50). After 21 days, seven of ten vaccinated rabbits must have survived, while controls
must have died from myxomatosis.
The manufacturer shall have established a minimum titre or potency taking into account loss in potency
during the shelf life.
2.
Method of manufacture
Shope’s fibroma virus is produced by multiple intradermal inoculations of seed virus into the skin on the back of a
number of rabbits. The product of fibroma homogenate can be stored by freezing or used immediately. Production
is also possible in rabbit dermal cell line. Only the second (and perhaps the third) viral passage can be used if
modification of the virus is to be avoided. After clarification by centrifugation, the supernatant fluid is mixed with a
stabiliser containing antibiotics and is distributed into ampoules or bottles for Iyophilisation. Kaolin may be added
as an adjuvant (40 mg/ml), in which case the vaccine is administered subcutaneously.
MV is produced in chicken embryo cells (derived from SPF eggs) or a suitable cell line, limiting the passage
number to a maximum of five. Virus is harvested after 2–6 days. The viral suspension may be stored at –70°C.
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The vaccine is prepared by diluting in specified proportions the viral preparation with a stabiliser for Iyophilisation.
After homogenisation, the product is distributed into bottles for Iyophilisation, the bottles being sealed under
vacuum or in sterile nitrogen.
Each virus can also be produced in RK-13 cells.
3.
In-process control
The Shope’s fibroma virus titre is measured by calculating the ID50 after intradermal inoculation of serial dilutions
of the clarified supernatant fluid into several sites (e.g. five) on up to six rabbits. A dilution of a standard
preparation of Shope’s fibroma virus is also inoculated into each rabbit to confirm the animal’s correct response to
inoculation. The titration can also be performed in a rabbit cell line. In each case the titre should correlate with the
required potency as defined by the test for efficacy, see Section C.1.c.
The identity of MV is checked in RK cells. Titration of each virus can also be done in RK-13 cells (TCID50).
Testing for contaminating viruses is done by inoculating a confluent monolayer of Vero cells. Vaccine, adjusted to
the equivalent of 20 doses/ml, is neutralised with an equal volume of monospecific hyperimmune serum for
30 minutes at 37°C. The mixture is filtered through a 0.22 µm membrane filter, and 1 ml volumes are inoculated
into five 25 ml bottles of cell cultures. These are kept under observation for 7 days. After harvesting, the cells are
suspended in medium and subjected to several freeze–thaw cycles, followed by centrifugation and filtration, and
the material is inoculated into fresh cultures and observed for 7 days. There should be no evidence of CPE or
haemadsorption to chicken RBCs.
4.
Batch control
a)
Sterility
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
b)
Safety
After rehydration, ten doses of the lyophilised fibroma vaccine are injected subcutaneously into each of three
susceptible rabbits, which are then observed for 21 days. Local reactions should be slight, with no
generalisation and no effect on general health.
Myxoma vaccine is tested using ten doses injected intradermally into the ears of three susceptible rabbits,
which are then observed for 21 days. The primary myxoma lesion should remain mild.
c)
Potency
The batch potency is determined by measurement of virus content. Serial dilutions of the vaccine are
inoculated into suitable cell cultures. One dose of vaccine shall contain not less than the minimum titre
established in Section C.1.c.
If the vaccine strain is not adapted to cultures, an efficacy test in rabbits shall be carried out (see Section C.1.c).
d)
Duration of immunity
Several groups of ten susceptible rabbits are vaccinated. One batch is tested by challenge infection (as in
the batch potency test), at 1, 2, 3, etc., months post-vaccination for Shope’s fibroma virus, and at 1, 3, 6, and
9 months for MV. The duration of immunity is deduced from the time during which at least seven of the ten
rabbits prove to be resistant to infection.
e)
Stability
Titrations of vaccine virus are carried out at intervals until 3 months beyond the requested shelf life on at
least three batches of vaccine.
5.
Tests on the final product
a)
Safety
See Section C.4.b.
b)
Potency
See Section C.4.c.
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REFERENCES
ARGUELLO VILLARES J.L. (1986). Contribución a la profilaxis de la mixomatosis del conejo mediante el uso de una
ceba homologa. Medicina Veterinaria, 3, 91–103.
ARTHUR C.P. & LOUZIS C. (1988). A review of myxomatosis among rabbits in France. Rev. sci. tech. Off. int. Epiz.,
7, 937–976.
BERTAGNOLI S., GELFI J., LEGALL G., BOILLETOT E., VAUTHEROT J.F., RASSCHAERT D., LAURENT S., PETIT F., BOUCRAUTBARALON C. & MILON A. (1996). Protection against myxomatosis and rabbit viral hemorrhagic disease with
recombinant myxoma virus expressing rabbit hemorrhagic disease virus capsid protein. J. Virol., 70, 5061–5066.
BRUN A., SAURAT P., GILBERT Y., GODART A. & BOUQUET J.F. (1981). Données actuelles sur l’épidémiologie, la
pathogénie et la symptomatologie de la myxomatose. Rev. Med. Vet., 132, 585–590.
CHANTAL J., BOUCRAUT-BARALON C., GANIERE J.P., PETIT F., PY R. & PICAVET D.P. (1993). Réaction de fixation du
complément en plaques de microtitration: application à la sérologie de la myxomatose. Etude comparative des
résultats avec la réaction d’immunofluorescence indirecte. Rev. sci. tech. Off. int. Epiz., 12, 895–907.
FENNER F. (1994). Myxoma virus. In: Virus Infections of Vertebrates, Vol. 5. Virus Infections of Rodents and
Lagomorphs, Osterhaus A.D.M.E., ed. Elsevier Science B.V., Amsterdam, Netherlands, 59–71.
FENNER F. & RATCLIFFE F.N. (1965). Myxomatosis. Cambridge University Press, London, UK.
FENNER F. & WOODROOFE G.M. (1954). Protection of laboratory rabbits against myxomatosis by vaccination with
fibroma virus. Aust. J. Exp. Biol., 32, 653–668.
GELFI J., CHANTAL J., PHONG T.T., PY R. & BOUCRAUT-BARALON C. (1999). Development of an ELISA for detection of
myxoma virus-specific rabbit antibodies; test evaluation for diagnostic applications on vaccinated and wild rabbit
sera. J. Vet. Diagn. Invest., 11, 240–245.
GILBERT Y., PICAVET D.P. & CHANTAL J. (1989). Diagnostic de la myxomatose: mise au point d’une technique
d’immunofluorescence indirecte. Utilisation de prélèvements sanguins sur papier buvard pour la recherche
d’anticorps. Rev. sci. tech. Off. int. Epiz. 8, 209–220.
GORSKI J. & MIZAK B. (1985). Polish vaccine against myxomatosis in rabbits. Med. Weter., 41, 113–116.
JOBERT R. (1983). Contribution à l’étude de la vaccination contre la myxomatose, vaccination à l’aide du virus
fibromateux. Thèse Doctorat Vétérinaire Toulouse n° 82.
JOUBERT L. (1973). La Myxomatose T.II. Série : Les Maladies Animales à Virus. L’Expansion éditeur, Paris,
France.
JOUBERT L., DUCLOS P.H. & TUAILLON P. (1982). La myxomatose des garennes dans le Sud-Est: la myxomatose
amyxomateuse. Rev. Med. Vet., 133, 739–753.
MARLIER D. & VINDEVOGEL H. (1996). La myxomatose amyxomateuse: isolement de trois souches en Belgique.
Ann. Med. Vet., 140, 343–346
RODAK L., SMID B., VALICEK L. & JURAK E. (1985). Collection of microvolume blood samples into glass capillaries for
the detection of antibody against Aujeszky’s disease virus in pigs by enzyme-linked immunosorbent assay
(ELISA) and solid-phase radioimmunoassay (RIA). Acta Vet. (Brno), 54, 207–216.
ROSELL J.M., GONZALES J.L., RUEDA A., GALLEGO E. & FLORES J.M. (1984). Mixomatosis atipica en Espana.
Medicina Veterinaria, 1, 401–412.
SAURAT P., CHANTAL J., GANIERE J.P., GILBERT Y., PICAVET D.P & LEFORT C. (1980). La réponse immunitaire dans la
myxomatose. Etude de la réponse humorale. Bull. Mens. Off. Nation. Chasse, 12, 297–309.
SAURAT P., GILBERT Y. & GANIERE J.P. (1978). Etude d’une souche de virus myxomateux modifié. Rev. Med. Vet.,
129, 415–451.
SHOPE R.E. (1932). A filtrable virus causing a tumor like condition in rabbits and its relationship to virus
myxomatosis. J. Exp. Med., 56, 803.
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SOBEY W.R., CONOLLY D. & ADAMS K.M. (1966). Myxomatosis: a simple method of sampling blood and testing for
circulating soluble antigens or antibodies to them. Aust. J. Sci., 28, No. 9, 354.
TOZZINI F. & MANI P. (1975). Studio su alcune carrateristiche di crescita del virus della mixomatosi ceppo Pisa 73.
Arch. Vet. Ital., 26, 19–26.
VON DER AHE C., DEDEK J. & LOEPELMANN H. (1981). Ergebnisse und Erfahrungen in der DDR bei der staatlichen
Prüfung und Praxiserpropung der aus der CSSR emportierten Myxomatose-Vakzine. Monatshe. Veterinarmed.,
36, 492–495.
*
* *
NB: There is an OIE Reference Laboratory for Myxomatosis
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for myxomatosis
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CHAPTER 2.6.2.
RABBIT HAEMORRHAGIC DISEASE
SUMMARY
Rabbit haemorrhagic disease (RHD) is a highly contagious and acute fatal disease of the European
rabbit (Oryctolagus cuniculus), caused by a calicivirus (genus Lagovirus). A similar disease, termed
European brown hare syndrome (EBHS), has been described in the hare (Lepus europaeus). Its
aetiological agent is a different species of lagovirus, antigenically related to the RHD virus (RHDV).
RHD is characterised by high morbidity and high mortality (70–90%), and spreads very rapidly by
direct and indirect transmission. Infection can occur by nasal, conjunctival or oral routes.
Transmission of RHD is facilitated by the high stability of the virus in the environment. The
incubation period varies from 1 to 3 days, and death usually occurs 12–36 hours after the onset of
fever. The clinical manifestations have been described mainly in the acute infection and are
nervous and respiratory signs, apathy and anorexia. Clear and specific lesions, both gross and
microscopic, are present. There is primary liver necrosis and a massive disseminated intravascular
coagulopathy in all organs and tissues. The most severe lesions are in the liver, trachea and lungs.
Petechiae are evident in almost all organs and are accompanied by poor blood coagulation. In
rabbits younger than 4–5 weeks, the RHDV infection course is subclinical.
Identification of the agent: The liver, spleen and blood of rabbits that died of acute RHD contain a
very high concentration of virus. Direct diagnosis is therefore easy and several tests can be used
However, as no sensitive cell substrates are established in-vitro isolation cannot be used. As a
consequence, the only available biological assay is reproduction of the disease in sensitive rabbits.
The main laboratory tests used are RNA amplification (reverse-transcription polymerase chain
reaction [RT-PCR]) and sandwich enzyme-linked immunosorbent assay (ELISA) based on the use
of monoclonal antibodies (MAbs). As RHDV haemagglutinates human Group O red blood cells very
efficiently, the haemagglutination (HA) test can also be used. False-negative results could arise in
the few rabbits that die from the chronic form of RHD (i.e. 4–8 days post-infection). In these rabbits
both an anti-RHDV IgM and virus-like particles (VLPs), made by the inner shell of RHDV, heavily
interfere with HA and some ELISAs. Moreover, using highly sensitive PCR in blood, intestinal
contents or faeces from healthy rabbits suspected to be RHD convalescent or RHDV re-infected,
some non-pathogenic RHDV-related caliciviruses were isolated and partially characterised;
importantly these RHDV-related viruses seem quite widespread in both commercial and wild rabbit
populations. The detection of calicivirus particles in liver homogenates or VLPs by electron
microscopy is also possible.
Serological tests: The level of RHDV-specific antibodies in the serum is a direct indication of the
state of protection of rabbits from RHD. In addition, determinating the levels for each class of
immunoglobulins (IgM, IgA and IgG) helps in distinguishing the first RHDV infection from reinfection or vaccination. Several types of ELISAs were developed and are currently used as a
replacement for the haemagglutination inhibition (HI) test initially used. The total level of RHDV-Ig is
semi-quantified using three main ELISAs: a) with the virus adsorbed on the solid phase; b) with the
virus and the serum both in liquid phase (competition ELISA), c) with the antigen (virus or subunits)
linked to the solid phase throughout MAbs. The different degree of exposition of the internal viral
epitopes obtained in these ELISAs helps to distinguish the Ig induced by RHDV from those induced
by the non-pathogenic related viruses.
Requirements for vaccines: Indirect control of the disease is achieved by vaccination using a
killed vaccine prepared from clarified liver suspensions of experimentally infected rabbits and
subsequently inactivated and adjuvanted. Vaccinated animals quickly produce solid protective
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Chapter 2.6.2. — Rabbit haemorrhagic disease
immunity against RHDV infection (within 7–10 days) and experimental data indicate that protection
lasts for a long period (over 1 year).
A. INTRODUCTION
Rabbit haemorrhagic disease (RHD) is a highly contagious and fatal disease of wild and domestic European
rabbits (Oryctolagus cuniculus) older than 2 months. RHD is caused by a calicivirus (genus Lagovirus, family
Caliciviriade), a non-enveloped small round RNA virus with only one major capsid protein (VP60) (Capucci et al.,
1990; Meyers et al., 1991; Ohlinger et al., 1990). The genus Lagovirus also includes the European brown hare
syndrome virus (EBHSV), the causative agent of a disease of brown hare (Lepus europaeus) termed EBHS, very
similar to RHD. Despite their high genetic relation (VP60 nucleotide similarity of 70%), RHDV and EBHSV are two
distinct viral species (Lavazza et al., 1996; Wirblich et al., 1994). There is only one serotype of RHDV and it
consists of two subtypes: RHDV (wild type) and RHDVa (Capucci et al., 1998; Schirrmeier et al., 1999).
RHD was first reported in 1984 in China (People’s Rep. of) (Lui et al., 1984) and 2 years later in Europe. To date,
RHD has been reported in over 40 countries in Asia, Africa, Americas, Europe and Oceania and is endemic in
most parts of the world (McIntosh et al., 2007). The RHDVa subtype has been identified in Europe since 1996–97
(Capucci et al., 1998; Schirrmeier et al., 1999) and is replacing the original RHDV in several parts of the world.
However, considering the RHDV genetic sequences deposited at the NCBI databank, the presence of RHDVa in
China (People’s Rep. of) dates back to 1985.
RHD is characterised by high morbidity and a mortality rate that usually ranges between 70 and 90%. The
incubation period varies from 1 to 3 days, and death usually occurs 12–36 hours after the onset of fever. In the
few rabbits that survive (5–10%), a specific and relevant IgM response appears within 3 days immediately
followed by an IgA and by an IgG response 2–3 days later (Barbieri et al., 1997). This antibody response triggers
the virus-clearance mechanism. In spite of this prompt humoral response, 5–10% of infected rabbits die between
4 and 8 days post-infection from a chronic form of RHD. In the liver and spleen of these rabbits, an RHD virus-like
particle (VLP) has been detected instead of RHDV (Barbieri et al., 1997; Capucci et al., 1991; Granzow et al.,
1996). This VLP is characterised by the lack of the outer shell on the viral capsid made up by the half C-terminal
portion of the VP60. As a consequence, this VLP is negative in the haemagglutination (HA) test as well as with
anti-RHDV monoclonal antibodies (MAbs) directed to outer conformational epitopes (Capucci et al., 1995). In the
blood and faeces of convalescent rabbits, as well as in rabbits infected with RHDV but already protected by
specific antibodies previously acquired (i.e. vaccinated or survivors of infection), the viral RNA is detected using
polymerase chain reaction (PCR) up to 15 weeks after the infection (Gall et al., 2007; Gall & Schirrmeier, 2006).
Whether this is the consequence of a slow viral clearance or of a real and prolonged virus replication
(persistence) is still to be established.
As a result of RHD serology testing (Capucci et al., 1991; 1997; Collins et al., 1995; Cooke et al., 2000), several
rabbit caliciviruses related to RHDV but non-pathogenic have been isolated and partially characterised in Europe
and Oceania (Capucci et al., 1996b; 1997; Forrester et al., 2002; Moss et al., 2002; Strive et al., 2009; White et
al., 2004). These caliciviruses induce a serological response that may interfere with and complicate RHD
serological diagnosis (Capucci et al., 1991; Cooke et al., 2000; Marchandeau et al., 2005; Nagesha et al., 2000;
Robinson et al., 2002).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
The liver contains the highest viral titre (from 103 LD50 [50% lethal dose] to 106.5 LD50/ml of 10% homogenate)
and is the organ of choice for viral identification for both RHDV and EBHSV. The amount of virus present in other
parts of the body is directly proportional to vascularisation; thus spleen and serum may serve as alternative
diagnostic materials, albeit suboptimal. Higher levels of VLPs have been reported in the spleen than in the liver of
animals that died from a subacute/chronic form of RHD (Barbieri et al., 1997; Capucci et al., 1991; Granzow et al.,
1996). The initial treatment of the diagnostic samples is almost identical irrespective of the diagnostic method to
be applied, with the exception of immunostaining techniques. An organ fragment is mechanically homogenised in
5–20% (w/v) phosphate buffered saline solution (PBS), pH 7.2–7.4, filtered through cheesecloth and clarified by
centrifugation at 5000 g for 15 minutes. At this stage, the supernatant can be directly examined by the HA test or
enzyme-linked immunosorbent assay (ELISA). If the sample is to be observed by electron microscopy (EM), it is
advisable to perform a second centrifugation at 12,000 g for 15 minutes, before the final ultracentrifugation. For
detection by PCR, viral RNA from the samples may be also directly extracted from tissues.
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a)
Enzyme-linked immunosorbent assay
Virus detection by ELISA relies on a ‘sandwich’ technique and several variations of this have been
described. One procedure uses the reagents, solutions, times and temperature that are used in the
competitive ELISA (C-ELISA) for serology (see Section B.2.b), except that the Tween 20 concentration is
twofold (0.1% [v/v]). The microplate used should be of high adsorption capability (e.g. Nunc Maxisorp
immunoplate). The liver homogenate is a 10% (w/v) suspension in standard PBS; 50 µl/well is the standard
volume to use in each step. The ELISA buffer used for all steps is PBS with 1% yeast extract (or bovine
serum albumin [BSA]), and 0.1% Tween 20, pH 7.4. All incubation steps are for 50–60 minutes at 37°C with
gentle agitation. After all steps three washes of 3–5 minutes must be performed using PBS with 0.05%
Tween 20. A positive and negative RHD rabbit liver homogenate must be used as controls. The horseradish
peroxidase (HRPO) conjugate could be purified lgG from a specific polyclonal serum or MAbs (see Section
B.2.b). Anti-RHDV MAbs have been produced in several laboratories and can be used instead of rabbit
polyclonal sera. MAbs recognising specific epitopes expressed only by the RHDVa variant were also
produced (Capucci, pers. data).
To better characterise the antigenicity of the RHD isolates by sandwich ELISA, it is advisable to test each
sample in four replicates, and then to use four different HRPO conjugates, i.e. two MAbs recognising the
same antigenic determinant present on the virus surface and expressed alternatively by the ‘classical’ strain
or by the RHDVa variant, a polyclonal hyperimmune anti-RHDV serum (which could identify potential ‘new
variant’ or correlated calicivirus, such as EBHSV) and a pool of MAbs recognising internal epitopes that can
detect smooth, degraded VLPs as well as EBHSV. An alternative antigen-capture ELISA has been
described using a sheep anti-RHDV as the capture antibody and a MAb for detection of RHDV (Collins et al.,
1996).
•
Test procedure (example)
For steps that are not specifically indicated see the procedure of the C-ELISA for serology (Section B.2.b).
i)
Coat the plate with anti-RHDV hyperimmune serum and the negative RHDV serum. The latter serves
as control for nonspecific reactions (false-positive samples). For each sample, four wells must be
sensitised with the positive serum and four wells with the negative one.
ii)
Dilute the liver extract to 1/5 and 1/30 (two replicates for each dilution) in ELISA buffer (see above),
directly in the wells of the plate (e.g. add 45 µl of the buffer into all the wells of the plate, add 10 µl of
the sample to the first two wells and then, after rocking, transfer 9 µl to the second wells). Treat the
controls, both positive and negative, in the same way as the samples.
iii)
After incubation and washing (see above), incubate with the HRPO conjugate.
iv)
After a last series of washing, add the chromogenic substrate. Orthophenylene-diamine (OPD) can be
used as peroxidase substrate for the final development of the reaction. Use 0.15 M citrate phosphate
buffer, pH 5.0, with 0.5 mg/ml OPD and 0.02% H2O2. The reaction is stopped after 5 minutes by the
addition of 50 µl of 1 M H2SO4.
v)
Absorbance is read at 492 nm. Positive samples are those showing a difference in absorbance >0.3,
between the wells coated with RHDV-positive serum and wells coated with the negative serum.
Usually, at the dilution 1/30, positive samples taken from rabbits with the classical acute form of RHD
give an absorbance value >0.8, while the absorbance value of the negative sample, at the dilution 1/5,
ranges from 0.1 to 0.25.
For diagnosis of EBHSV, it is possible to use this RHDV-specific sandwich ELISA, but, due to the high
antigenic difference existing between the two viruses, there is a risk of obtaining false-negative results.
Therefore, the adoption of an EBHSV-specific sandwich ELISA technique using either a high-titre positive
anti-EBHSV hare serum, or cross-reacting RHDV MAbs (Capucci et al., 1991; 1995), or specific EBHSV
MAbs, instead of rabbit serum, is highly recommended (Capucci et al., 1991).
b)
Nucleic acid recognition methods
The application of the RT-PCR to the detection of RHDV-specific nucleic acid has been described by several
authors (Gould et al., 1997; Guittre et al., 1995). Owing to the low level of sequence variation among RHDV
isolates and the high sensitivity of PCR, reverse transcription (RT)-PCR represents an ideal rapid diagnostic
test for RHD. This method is carried out on organ specimens (optimally liver), urine, faeces and sera using
different oligonucleotide primers derived from the capsid region of the RHDV genome (N-terminal portion).
The RHD OIE Reference Laboratory uses a single-step RT-PCR with the following primers specific for the
PV60 gene: [forward: 5’-CCT-GTT-ACC-ATC-ACC-ATG-CC-3]’, [reverse: 5’-AAC-CCT-CCA-GGT-ACTGGT-TG-3’], whereas Guittre et al. used [forward: 5’-GAG-CTC-GAG-CGA-CAA-CAG-GC-3]’, [reverse: 5’CAA-ACA-CCT-GAC-CCG-CGA-AC-3’]. cDNA obtained from the RT reaction is usually PCR amplified as
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Chapter 2.6.2. — Rabbit haemorrhagic disease
described (Guittre et al., 1995) or using one step standard RT-PCR methodology. To reveal the PCR
product, the amplified DNA reaction mixture is subjected to electrophoresis on agarose gel. If needed,
specificity of the PCR product can be determined by sequencing or by Southern blot and hybridisation with a
radioactively labelled internal probe. A similar RT-PCR method has been used to identify the nonpathogenic
RCV (Capucci et al., 1998). Several primers, specific for the RHDV RNA polymerase gene and
complementary to the VP60 and ORF2 genes, are used and the amplified fragments are subjected to
Southern blot analysis. RT-PCR represents an extremely sensitive method for the detection of RHDV, and is
104-fold more sensitive than ELISA (Guittre et al., 1995). It is not strictly necessary for routine diagnosis, but
it is more sensitive, convenient and rapid than other tests.
Similarly an RT-PCR for the detection of EBHSV has been applied to the detection and characterisation of
EBHSV stains (Le Gall-Reculé et al., 2001]).
An internally controlled multiplex real-time RT-PCR using TaqMan® fluorogenic probes and external
standards for absolute RNA quantification has been developed recently as a further diagnostic tool for the
detection of RHDV. The test revealed a specificity of 100%, an analytical sensitivity of 10 copies/well and a
linearity over a range from 101 to 1010 copies. The method has been used to quantify RHDV RNA in
experimental infection of vaccinated rabbits and in RHD convalescent rabbits (Gall et al., 2007; Gall &
Schirrmeier, 2006). The oligonucletides used in this method are: [VP60-7_forward: 5’-ACY-TCA-CTG-AAC
TYA-TTG-ACG-3’, vp60-8_reverse: 5’ TCA-GAC-ATA-AGA-AAA-GCC-ATT-GG-3’] and probe [VP60-9_fam
5’-FAM-CCA-ARA-GCA-CRC-TCG-TGT-TCA-ACC-T-TAMRA-3’].
An in-situ hybridisation technique using either sense or antisense DNA probes has been developed for
investigating the presence of RHDV in tissue samples (Gelmetti et al., 1998). This method is highly sensitive
and can be used for early diagnosis of RHD as it gives positive results as soon as 6–8 hours post-infection.
However, it is expensive and difficult to carry out, and thus it is mainly indicated for research studies.
c)
Electron microscopy
Negative-staining EM can be performed using the so-called ‘drop method’. A formvar/carbon-coated grid is
placed on a drop of organ suspension (prepared as described in Section B.1), and left for 5 minutes. After
removing excess fluid with the edge of a torn piece of filter paper, the grid is put to float on a drop of 2%
sodium phosphotungstate (NaPT), pH 6.8, for 1.5 minutes. Excess stain is removed and the grid is finally
observed at ×25,000 magnification.
Due to the lower sensitivity of the drop method, it is advisable to ultracentrifuge the sample in order to
concentrate the viral particles. The pellet obtained after ultracentrifugation (at least 100,000 g for 30 minutes
or, alternatively, using Beckman Airfuge at 21 psi for 5 minutes) is resuspended in PBS or distilled water, put
on to a grid for a few minutes, and then stained as described. RHD virions are visible as uncoated particles,
32–35 nm in diameter, presenting an inner shell (25–27 nm in diameter), delineated by a rim from which
radiate ten short regularly distributed peripheral projections. Smooth (s-RHDV) particles are identified by the
complete loss of external portions, becoming perfectly hexagonal and smaller, with only the capsid rim
visible (Barbieri et al., 1997; Capucci et al., 1991; Granzow et al., 1996).
For diagnostic purposes and especially when other methods give doubtful results, the best EM method is an
immuno-EM technique (IEM). This method uses either a hyperimmune anti-RHDV serum, obtained from
rabbit or other species, or specific MAbs, which are incubated with an equal volume of the sample for 1 hour
at 37°C before ultracentrifugation. The immunological reaction induces the clumping of the viral particles into
aggregates that are quickly and easily identified by EM. Immunogold methods can also be applied to better
identify virions and viral proteins.
EBHSV can also be identified in diagnostic samples by EM examination. In addition, the IEM method using
convalescent anti-EBHSV serum or specific anti-EBHS MAbs can be used to identify EBHSV. By using
antisera that is specific for EBHSV and RHDV, it is possible to differentiate between the two viruses.
d)
Haemagglutination test
HA was the first test to be used for routine laboratory diagnosis of RHD (Lui et al., 1984|]). It should be
performed with human Group O red blood cells (RBCs), freshly collected, stored overnight in Alsever’s
solution, and washed in 0.85% PBS at pH 6.5 (range 6–7.2). HA is less evident or non-existent when RBCs
of other species are used. Washed RBCs are suspended at 0.75% in PBS. A twofold dilution of the clarified
supernatant of a 10% tissue homogenate of liver or spleen is incubated with an equal volume of washed
RBCs in a sealed round-bottom microtitre plate at, preferably, 4°C. After 1 hour (range from 20 minutes to
2 hours) of incubation, agglutination at an end-point dilution of >1/160 is considered to be positive. Lower
titres should be regarded as suspicious, and should be checked using other methods. Around 10% of
samples found to be positive by ELISA or EM give negative results in HA (HA false-negative). Some RHD
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isolates may exhibit temperature-dependent differences in haemagglutinating characteristics (Capucci et al.,
1996a) and could show HA activity only when the test is performed at 4°C. Nevertheless, the HA false
negativity is mainly detected in organs of rabbits showing a subacute/chronic form of the disease and it
depends on the characteristics of the VLPs.
Hare organs rarely give a significant titre when the RHDV HA protocol is used. To demonstrate HA activity in
organs from EBHSV-infected rabbits, a modified procedure should be adopted: all steps are carried out at
4°C, the organ suspension is treated with an equal volume of chloroform, and RBCs are used at a pH not
higher than 6.5 (Capucci et al., 1991). Even using this method, only about 50% of the samples give positive
results. This is because the disease of hares is often subacute or chronic and therefore the virus has the
antigenic and structural characteristics typical of the VLPs particles (Capucci et al., 1991).
Because of the practical difficulty of obtaining and keeping and the risk from working with human Group O
blood cells, and because of the difficulty of obtaining consistent results, this test has been replaced by the
virus-detection ELISA.
e)
Immunostaining
Tissue fixed in 10% buffered formalin and embedded in paraffin can be immunostained using an avidin–
biotin complex (ABC) peroxidase method. The sections are first deparaffinised in xylene and alcohol,
counter-stained with haematoxylin for 1 minute and rinsed in tap water. They are then put in a methanol bath
containing 3% H2O2 and washed in PBS three times for 5 minutes each. To limit background interference
caused by nonspecific antibody binding, the samples are incubated with normal rabbit serum for 1 hour at
room temperature prior to the addition of biotin. The slides are incubated overnight in a humid chamber at
room temperature with biotinylated rabbit anti-RHDV serum or MAbs, are washed as before and incubated
again for 30 minutes at 37°C with an ABC peroxidase. The slides are then washed three times. Amino-ethylcarbazole is used as substrate. Finally, the slides are rinsed in tap water and mounted (Stoerckle-Berger et
al., 1992).
Intense nuclear staining and diffuse cytoplasmic staining of necrotic cells in the liver, mainly in the periportal
areas, are characteristic and specific. Positive staining of macrophages and Kupffer’s cells is also observed,
as well as hepatocellular reactions. Positive reactions can also be detected in the macrophages of the lungs,
spleen and lymph nodes, and in renal mesangial cells (Stoerckle-Berger et al., 1992).
Tissue cryosections fixed in methanol or acetone can be directly immunostained by incubation for 1 hour
with fluorescein-conjugated rabbit anti-RHDV serum or MAbs. Specific fluorescence can be detected in the
liver, spleen, and renal glomeruli.
f)
Western blotting
When other tests such as HA or ELISA give doubtful results (low positivity) or the samples are suspected of
containing s-RHDV particles, western blotting analysis is useful for determining the final diagnosis.
Homogenates are prepared as described previously, and virus particles are further concentrated (tenfold) by
ultracentrifugation (100,000 g for 90 minutes) through a 20% (w/w) sucrose cushion.
Both the supernatant and the pellet can be examined to detect, respectively, the RHDV 6S subunits
(Capucci et al., 1995) and the denatured VP60 structural protein of RHDV or its proteolytic fragments, which
can range in size from 50 to 28 kDa. A positive and negative control samples should be used on each
occasion.
RHDV proteins could be detected with polyclonal antibodies or MAbs. If MAbs are used, they should
recognise continuous epitopes. RHDV-specific MAbs recognising internal or buried epitopes could be used
also to detect EBHSV. Rabbit anti-RHDV hyperimmune sera are less efficient than MAbs at recognising the
same band patterns (Capucci et al., 1996b).
Sample proteins are denatured for 2 minutes at 100°C in the presence of 60 mM Tris, pH 6.8, 2% sodium
dodecyl sulphate (SDS), 2% beta-mercaptoethanol, and 5% glycerol, separated on 10% SDS/PAGE
(polyacrylamide gel electrophoresis), and then transferred by electroblotting to nitrocellulose or PVDF
(polyvinylidene flouride) membranes, in 25 mM Tris, 192 mM glycine pH 8.3 and 20% (v/v) methanol at 1.5 Å
for 60 minutes with cooling or at 0.15 A overnight. After transfer the membranes are saturated for 30–
60 minutes in blocking buffer or PBS, pH 7.4 containing 2% bovine serum albumin (BSA), and subsequently
incubated for 2 hours at room temperature with the appropriate serum dilution in PBS, pH 7.4, and 1% BSA.
The filters are washed thoroughly with PBS and incubated for 1 hour at room temperature with anti-species
alkaline phosphatase-labelled immunoglobulins at a dilution predetermined by titration. Finally, the filters are
again washed and the chromogenic substrate (5-bromo-4-chloro-3-indolylphosphate nitro blue tetrazolium)
is added.
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Positive test samples and the positive control will produce a pattern consistent with reaction to proteins of
molecular weights of, respectively, 60 kDa (the single structural protein of RHDV) or 41–28 kDa (the
fragments of the VP60 associated with the transition from RHDV to s-RHDV), when examining the pellet,
and 6 kDa (the subunits) when examining the supernatant.
Western blot analysis can also be used to identify EBHSV. The test procedure is identical. The pattern of
protein bands, detected using either an anti-EBHSV polyclonal serum or cross-reacting anti-RHDV MAbs, is
similar. However, the percentage of samples showing viral degradation is higher and therefore several
fragments of lower molecular weight, originating from the VP60 structural protein, are often observed.
g)
Rabbit inoculation
As no efficient in-vitro replication system has been established for RHDV and EBHSV, cell culture isolation
cannot be included among the diagnostic methods. Rabbit inoculation therefore remains the only way of
isolating, propagating and titrating the infectivity of the RHDV. However, experimental infection of rabbits is
not a practical method for the routine diagnosis of RHD and should be considered only when all other
methods give inconclusive results. When this occurs, the rabbits involved must be fully susceptible to the
virus, i.e. they should be over 2 months old and have no RHDV antibodies (see serological methods). RHD
can be reproduced by using filtered and antibiotic-treated liver suspensions, inoculated either by the
intramuscular, intravenous or oro-nasal route. When the disease is clinically evident, the signs and postmortem lesions are similar to those described after natural infection. A rise in body temperature is registered
between 18 and 24 hours post-infection, followed, in 70–90% of cases, by death between Robinson et al.,
2002 and 72 hours post-infection. A few individuals may survive until 6–8 days after infection. Animals that
overcome the disease show only a transient hyperthermia, depression and anorexia, but present a striking
seroconversion that can be detected easily 3–4 days post-infection.
2.
Serological tests
Infection by RHDV can be diagnosed through detection of a specific antibody response. As the humoral response
has great importance in protecting animals from RHD, determination of the specific antibody titre after vaccination
or in convalescent animals is predictive of the ability of rabbits to resist RHDV infection.
Three basic techniques are applied for the serological diagnosis of RHDV: haemagglutination inhibition (HI) (Lui
et al., 1984), indirect ELISA (I-ELISA) and C-ELISA (Capucci et al., 1991). Each of these methods has
advantages and disadvantages. With respect to the availability of reagents and the technical complexity of
carrying out the test, HI is the most convenient method, followed by the I-ELISA and C-ELISA, respectively. On
the other hand, both ELISAs are quicker and easier than HI, particularly when a large number of samples are
tested. The specificity of the C-ELISA is markedly higher than those achieved with the other two methods
(Capucci et al., 1991). An alternative C-ELISA method has been described (Collins et al., 1995). For improved
serological interpretation and for correctly classifying the immunological status of rabbits, a combination of ELISA
techniques that distinguish IgA, IgM and IgG antibody responses is also available.
Some other additional tests (Capucci, unpublished data; Cooke et al., 2000) could be used for particular
investigations and when a higher level of sensitivity is needed to detect antibodies in non-target species or
antibodies induced by cross-reacting RHDV-like agents (see Section A. Introduction).
They are:

I-ELISA: the antigen is linked to the solid phase by a RHDV-specific MAb (1H8). It has a slightly higher
sensitivity than C-ELISA, making possible measurement of highly cross-reactive antibodies and it can detect
antibodies with low avidity.

Solid-phase ELISA (SP-ELISA): the purified antigen is directly adsorbed to the solid phase and because of
virus deformation, internal epitopes are exposed. Therefore it detects a wider spectrum of antibodies and
has high sensitivity and low specificity. For these reasons it can also be used for EBHSV serology.

Sandwich ELISA to detect IgM and IgG in liver or spleen samples already examined with the virological test:
such a test is particularly useful in those animals that die from the ‘chronic’ form of the disease, when
detection of the virus may be difficult. In this case, a high level of RHDV-specific IgM and a low level, if any,
of IgG are the unambiguous markers of positivity for RHD.
a)
Haemagglutination inhibition
Antigen: The antigen is prepared using infected rabbit liver collected freshly at death. The liver is
homogenised in 10% (w/v) PBS, pH 6.4, and clarified by two consecutive low speed centrifugations (500 g
for 20 minutes and 6000 g for 30 minutes). The supernatant, drawn from the tube so as to avoid the
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superficial lipid layer, is filtered through a 0.22 µm pore size mesh, titrated by HA, and divided into aliquots,
which are stored at –70°C.
Serum samples: Before testing, sera are inactivated by incubation at 56°C for 30 minutes. The sera are then
treated with 25% kaolin (serum final dilution: 1/10) at 25°C for 20 minutes and centrifuged. This is followed
by a second kaolin treatment, also at 25°C for 20 minutes, this time with 1/10 volume of approximately 50%
packed human Group O RBCs. These are freshly collected, stored overnight in Alsever’s solution and
washed in 0.85% PBS, pH 6.5. The sera are clarified by centrifugation.

Test procedure
i)
Dispense 50 µl of serum into the first well of a round-bottom microtitre plate and make double dilutions
into wells 2–8 using PBS with 0.05% BSA.
ii)
Add 25 µl of RHDV antigen containing 8 HA units to each well and incubate the plate at 25°C for 30–
60 minutes.
iii)
Add 25 µl of human Group O RBCs at 2–3% concentration to each well and allow to settle at 25°C for
30–60 minutes.
iv)
Titrate the antigen with each test to ensure that 8 HA/ 25 µl were used, and include positive and
negative serum controls.
The serum titre is the end-point dilution showing inhibition of HA. The positive threshold of serum titres is
correlated to the titre of the negative control sera; it usually is in the range 1/20–1/80.
Because of the practical difficulty of obtaining, keeping and the risk from working with human Group O blood
cells, and because of the difficulty in obtaining consistent results, this test is being superseded by the
serological or antibody-detection ELISA.
b)
Competitive enzyme-linked immunosorbent assay
Antigen: An international standard strain is not yet available; however, as only one serotype has been
identified so far world-wide, reliable results can be obtained by different laboratories each using their own
standard virus. Even the antibodies induced by the identified RHDV variants are recognised by the standard
method described here. In addition, the test can also easily detect antibodies originating from infection of
rabbits with the nonpathogenic RCV, because of its high genetic correlation with RHDV (Capucci et al.,
1996b; 1997).
The antigen can be prepared as described previously for HI (Section B.2.a), taking care to store it at –20°C
in the presence of glycerol at 50% (v/v) to prevent freezing. If necessary, the virus can be inactivated before
the addition of glycerol, using 1.0% binary ethylenimine (BEI) at 33°C for 24 hours. Antigen must be
pretitrated in ELISA and then used as the limiting reagent: i.e. the dilution that corresponds to 60–70% of the
plateau height (absorbance value at 492 nm in the range 1.1–1.3).
Anti-RHDV serum: specific polyclonal sera with high anti-RHDV titre can be obtained in different ways. Two
possible and currently used methods are as follows:
i)
Rabbits are infected with a RHDV-positive 10% liver extract diluted 1/100 in PBS to obtain
convalescent sera (21–25 days) containing a high level of anti-RHDV IgG. Due to the high mortality
rate associated with RHDV, it is necessary to infect at least 10–15 seronegative rabbits or to infect
rabbits that are only partially protected (e.g. 4–8 rabbits infected from 3 to 7 days post-vaccination).
Rabbits that survive the infection must be bled 21–25 days post-infection to obtain the convalescent
sera. Alternatively, convalescent rabbits can be re-infected 3–4 months post-infection and bled 10–
15 days later to obtain RHDV hyperimmune sera.
ii)
RHDV is purified from the livers of experimentally infected rabbits that died from an acute form of the
disease (between 28 and 40 hours post-Infection), using one of the methods that has been published
(Capucci et al., 1990; 1991; 1995; Meyers et al., 1991; Ohlinger et al., 1990). Then the purified RHDV
can be used to immunise sheep or goats according to classical protocols using oil adjuvants. The same
procedure can also be used to inoculate rabbits if the purified virus is inactivated before inoculation.
Anti-RHDV MAbs may be used instead of rabbit polyclonal sera. Purification of rabbit IgG and conjugation to
HRPO can be done following the standard protocols. The conjugated antibody is titred in a sandwich ELISA
in the presence and absence of RHDV antigen (negative rabbit liver). It is then used at the highest dilution
showing maximum (plateau high) absorbance (if the serum had a good anti-RHDV titre, the value of the
HRPO conjugate should range from 1/1000 to 1/3000).
Control sera: negative serum is taken from rabbits fully susceptible to RHDV infection. Positive serum is
either a convalescent serum diluted 1/100 in a negative serum or a serum taken from a vaccinated animal.
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
Test procedure (example)
i)
The rabbit anti-RHDV serum diluted to a predetermined titre, e.g. 1/5000 in 0.05 M carbonate/
bicarbonate buffer, pH 9.6, should be adsorbed to an ELISA microplate of high adsorption capability
(e.g. Nunc Maxisorb Immunoplate) at 4°C overnight.
ii)
Wash the plate three times for 3–5 minutes each time, in PBS, pH 7.4, with 0.05% Tween 20 (PBST).
When the plates are not immediately used, they can be stored, closed in a plastic bag, for 1 month at
–20°C.
iii)
Distribute 25 µl/well PBST with 1% yeast extract (PBSTY) or 1% BSA (PBST-BSA) to all the wells
needed on the plate (see below). Add 7 µl of the first serum sample to the first two wells (A1 and B1),
7 µl of the second serum to the second two wells (C1 and D1), and continue with the third (E1 and F1)
and the fourth (G1 and H1) sera, thus completing the first column. If qualitative data (positive/negative)
are needed, repeat the operation in the second column with sera samples from 5 to 8, and in the third
column with sera samples from 9 to 12, and so on. If the titre of the serum needs to be determined, the
serum must be diluted further. Agitate the plate and then use an eight-channel micropipette to transfer
7 µl from the wells in column 1 to the wells in column 2. This corresponds to a four-fold dilution of the
sera. This last operation can be repeated once (titre 1/160), twice (titre 1/640), or four times (titre
1/10,240). Either in the case of testing sera for qualitative data (single dilution), or for getting the final
titre (several dilutions), complete each plate leaving 12 wells free for the control sera. Add 7 µl of
positive sera to wells G7 and H7, and 7 µl of negative sera to wells G10 and H10, then dilute them
once and twice (1/40–1/160).
iv)
Add 25 µl/well antigen suspended in PBSTY to all the wells on the plate, at a dilution that is double the
decided dilution, as described above in the antigen section (see the first part of this ELISA method
description).
v)
Incubate the plate at 37°C on a rocking platform for 50–60 minutes.
vi)
Wash the plate as described in step ii.
vii)
Add 50 µl/well rabbit IgG anti-RHDV conjugated with HRPO at the decided dilution, as described above
in the ‘anti-RHDV serum’ section (see the first part of this ELISA test description).
viii) Incubate the plate at 37°C on a rocking platform for 50–60 minutes, and wash as described in step ii
adding a fourth wash of 3 minutes duration.
ix)
Use 50 µl/well OPD as hydrogen donor under the following conditions: 0.5 mg/ml OPD in 0.15 M
phosphate/citrate buffer, pH 5, and 0.02% H2O2. Stop the reaction after 5 minutes by addition of
50 µl/well 1 M H2SO4.
x)
Read the plate on a spectrophotometer using a 492 nm filter.
The serum is considered to be negative when the absorbance value of the first dilution (1/10) decreases by
less than 15% of the reference value (dilution 1/10 of the negative control serum), while it is positive when
the absorbance value decreases by 25% or more. When the absorbance value of the 1/10 dilution
decreases by between 15% and 25% of the reference value, the sera is considered to be doubtful.
The serum titre corresponds to the dilution giving an absorbance value equal to 50% (±10) of the average
value of the three negative serum dilutions.
A wide range of titres will be found, depending on the origin of the sample. Positive sera range from 1/640 to
1/10,240 in convalescent rabbits, from 1/80 to 1/640 in vaccinated rabbits and from 1/10 to 1/160 in ‘nonpathogenic’ infection. Knowing the origin of the sample allows a choice to be made between testing one or
more dilutions. Testing only the first dilution gives a positive or negative result. The titre is established by
testing all dilutions, up to the sixth one.
Due to the significant antigenic differences existing between RHDV and EBHSV (Capucci et al., 1990;
Stoerckle-Berger et al., 1992), the serological techniques described above, which use RHDV as antigen, are
not recommended for the serological diagnosis of EBHS. However, a direct ELISA method could be
employed for the detection of positive and negative EBHSV hare sera; in fact, the adsorption of RHDV on to
the solid phase of an ELISA microplate exposes cross-reactive antigenic determinants. Alternatively, a
specific C-ELISA for EBHSV can be arranged in a similar way, using specific reagent (antigen and antisera)
prepared as described above for RHDV.
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c)
Isotype enzyme-linked immunosorbent assays (isoELISAs)
These isoELISAs enable the detection and titration of isotypes IgA, IgM and IgG (Capucci et al., 1997).
Isotype titres are critical for the interpretation of field serology in four main areas: cross-reactive antibodies,
natural resilience of young rabbits, maternal antibodies, antibodies in previously infected rabbits (Cooke et
al., 2000). In fact in the case of passive antibodies, only IgG are detected; in vaccinated animals, no IgA are
detected and in recently infected rabbits, first IgM and then IgA and IgG are detected (Cooke et al., 2000).
To detect RHDV-specific IgG, one RHDV-specific MAb is adsorbed to the Maxisorp plate at a concentration
of 2 µg/mI by the method described above for the polyclonal serum in the C-ELISA (see above Section
B.2.b, test procedure step i). Virus is added to the plates at a concentration double that used in the C-ELISA
and after incubation and washing; sera are added and serially diluted four-fold starting from 1/40. A MAb
anti-rabbit IgG HRPO conjugate is used to detect IgG bound to the virus. The final step for the isoELISAs for
IgG, lgM and IgA is the addition of OPD and H2SO4 as for the C-ELISA. To detect IgM and IgA isotypes the
phases of the ELISA reaction are inverted in order to avoid competition with IgG, which is usually the
predominant isotype. MAb anti-rabbit IgM or anti-rabbit IgA is adsorbed to the wells and then the sera are
diluted as described above. Incubation with the antigen follows and then HRPO-conjugated MAb is used to
detect the RHDV bound to the plate. Sera are considered to be positive if the OD492 (optical density) value
at the 1/40 dilution is more than 0.2 OD units (two standard deviations) above the value of the negative
serum used as a control. The titre of each serum is taken as the last dilution giving a positive value. Because
isoELISA tests do not follow identical methodology, equivalent titres do not imply that isotypes are present in
the same amounts.
C. REQUIREMENTS FOR VACCINES
1.
Background
a)
Rationale and intended use of the product
In countries where RHD is endemic, indirect control of the disease in farmed animals and pet rabbits is
achieved by vaccination using the appropriate type of vaccine – one that is prepared from clarified liver
suspension of experimentally infected rabbits, and that is subsequently inactivated and adjuvanted. The
methods of inactivation (formaldehyde, beta-propiolactone or other substances) and the adjuvants used
(incomplete mineral oil or aluminium hydroxide), can vary according to the protocol used by the different
manufacturers.
Most vaccine manufacturers recommend a single basic vaccination, with yearly booster. Normally, a 1-ml
dose is inoculated subcutaneously in the neck region, or intramuscularly. The first injection should be given
at 2–3 months. In those units with no history of disease, where the anamnesis for RHD is negative, it is
advisable to vaccinate only the breeding stock. Considering the high restocking rate in industrial rabbit
farms, the usual vaccination programme is to administer the vaccine to all breeders, independently of their
age, every 6 months. This should ensure that all animals get at least one vaccination per year. Booster
vaccination is strongly recommended to ensure a good level of protection, although experimental data
indicate that protection usually lasts for a long time (over 1 year).
Given the short life-cycle (approximately 80 days) of fattening rabbits and their natural resistance to the
disease until 35–40 days of age, vaccinating these rabbits is not necessary if the situation on the farm is
normal, i.e. good biosecurity measures are applied and there are no outbreaks of the disease in the area.
Following an outbreak of RHD, even if strict hygiene and sanitary measures are adopted, including cleaning
and disinfection, safe disposal of carcasses and an interval before restocking, it is strongly recommended to
vaccinate meat animals at the age of 40 days, because the incidence of re-infection is very high. Only after
several production cycles is it advisable to stop vaccination of meat animals. To verify the persistence of
infective RHD inside the unit, a variable number of rabbits, starting with a small sentinel group, should not be
vaccinated.
Given that immunity starts after about 7–10 days, vaccination could also be considered a quite effective
post-exposure treatment. In some situations in particular it may be included in the emergency strategies
applied when RHD occurs on those farms having separate sheds and where good biosecurity measures are
regularly applied. Indeed, better results in limiting the diffusion of the disease and reducing economic losses
could be obtained by using seroterapy through the parenteral administration of anti-RHDV hyperimmune
sera, which produces a rapid, but short-lived, protection against RHDV infection.
Vaccine should be stored at 2–8°C and it should not be frozen, or exposed to bright light or high
temperatures.
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2.
Outline of production and minimum requirements for conventional vaccines
a)
Characteristics of the seed
i)
Biological characteristics
At present, RHDV replication can be obtained exclusively in susceptible animals. Therefore, the source
of seed virus for the production of inactivated tissue vaccines is infected liver homogenates obtained by
serial passages in rabbits that have been inoculated with a partially purified RHD viral suspension. The
rabbits used for inoculation are selected from colonies shown to be healthy and susceptible to the
disease by periodic serological testing.
ii)
Quality criteria (sterility, purity, freedom from extraneous agents)
The partially purified RHD viral suspension is obtained by centrifuging the 1/5 liver suspension (w/v) in
PBS at 10,000 g for 20 minutes at 4°C. The resulting supernatant is treated with 8% (v/v) polyethylene
glycol (PEG 6000) overnight at 4°C. The pellet is resuspended at a dilution of 1/10 in PBS, and
subsequently centrifuged at 10,000 g for 20 minutes at 4°C. The supernatant is ultracentrifuged at
80,000 g for 2 hours at 4°C through a 20% cushion of sucrose. The pellet is resuspended in PBS
(1/100 of the starting volume).
This viral suspension is then characterised by negative-stain EM examination, determination of
reactivity in ELISA, and capability of HA at room temperature (HA titre against RBCs of human Group
O higher than 1/1280).
The absence of viable bacteria, fungi or mycoplasma should be determined by using common
laboratory bacteriological methods. PCR methods may be used for the detection of extraneous viruses
(e.g. Myxoma virus [MV]).
Seed virus is controlled by direct inoculation into susceptible rabbits followed by evaluation of the
clinical signs in the course of the experimental infection. Suitable seed virus should cause the death of
70–80% of the rabbits within 24–72 hours post-inoculation, with the internal organ lesions characteristic
of RHD. To validate the test, gross and histopathological examination of all rabbits should be
performed to exclude undercurrent diseases.
Seed virus is titrated before use and should contain at least 105 LD50. It should be stored frozen
(–70°C), better with the addition of 1:1 volume of glycerol or freeze-dried.
b)
Method of manufacture
i)
Procedure
Following inoculation of susceptible rabbits, the liver and spleen of those rabbits that die between
24 and 72 hours post-inoculation are collected. Rabbits that died later must be discarded. The organs
are minced in 1/10 (w/v) sterile PBS, pH 7.2–7.4, and the mixture is homogenised for 10 minutes in a
blender in a refrigerated environment. The mixture is then treated with 2% chloroform (18 hours at
4°C), followed by centrifugation at 6000 g for 1 hour at 4°C. The supernatant is collected by high
pressure continuous pumping and is subsequently inactivated. The viral suspension is assayed by HA
test and ELISA and, once the number of HA units from the initial titration is known, more sterile PBS is
added in sufficient volume to provide, after inactivation and adsorption/addition of the adjuvant , a
concentration of 640–1280 HA units/ ml in the commercial product. Various agents have proved
effective at abolishing viral infectivity. The most frequently used are formaldehyde and betapropiolactone, which can be used at different concentrations and temperatures, for variable periods of
time and also in combination. During inactivation, it is advisable to continuously agitate the fluid.
Aluminium hydroxide, Freund’s incomplete adjuvant or another oil emulsion is then incorporated into
the vaccine as adjuvant. A preservative, thiomersal (merthiolate), is finally added at a dilution of
1/10,000 (v/v) before distribution into bottles.
ii)
Requirements for substrates and media
As the virus cannot be grown in vitro, the only requirements are those concerning infected animals.
Rabbits must be free from RHDV and myxomatosis virus and should not have anti-RHDV antibodies,
including cross-reactive antibodies induced by the non-pathogenic RHDV-related rabbit calicivirus
(RCV).
The animals (at least 4 months old) must be kept in strict quarantine on arrival, in a separate area and
reared under satisfactory health conditions (see Laboratory animal facilities in Chapter 1.1.3 Biosafety
and biosecurity in the veterinary microbiology laboratory and animal facilities).
Seed virus propagation and production of vaccine batches rely on the same protocol of experimental
infection, involving intramuscular injection of a dose of at least 100 LD50.
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iii)
In-process controls
Antigen content
The RHD titre is determined before inactivation by calculating the HA titre, which should be higher than
1/1280, and the ELISA reactivity. Both values are again determined after inactivation and
adsorption/addition of the adjuvant. Negative-staining EM confirms the identity of RHD.
Sterility
The organs are tested for the presence of viable bacteria, viruses, fungi or mycoplasma according to
the protocol used for testing master seed virus. PBS solution and aluminium hydroxide gel are
sterilised by autoclaving; oil emulsion is sterilised by heating at 160°C for 1 hour.
Inactivation
Before incorporation of the adjuvant, the inactivating agent and the inactivation process must be shown
to inactivate the vaccine virus under the conditions of manufacture. Thus, a test is carried out on each
batch of the bulk harvest as well as on the final product.
Thirty adult rabbits (>4 months of age) are used in three groups of 10. The first and second group are
injected with concentrated antigen and kept under observation for 15 and 7 days, respectively. The
second group is humanely killed after 7 days. The third group is injected with the liver of rabbits from
the second group and kept under observation for 21 days. The dose of the inoculum, administered
parenterally (intramuscular or subcutaneous), is 1 ml of concentrated antigen (PEG precipitation)
corresponding to at least 10 doses (HA ≥20480). The observation period is: 10 rabbits for 7 days,
10 rabbits for 15 days and 10 rabbits for 21 days. All the rabbits kept under observation must survive
without any clinical signs. The liver should give negative results using the HA test and sandwich ELISA.
The rabbits inoculated with antigen should have a positive serological titre (e.g. >1/80 using
competitive ELISA) and those injected with livers obtained after the first passage should be
serologically negative.
iv)
Final product batch tests
Sterility, safety and potency tests should be carried out on each batch of final vaccine; tests for
duration of immunity should be carried out once using a typical batch of vaccine, and stability tests
should be carried out on three batches.
Sterility/purity
Each batch of vaccine must be tested for the presence of viable bacteria, viruses, fungi or mycoplasma
according to the same protocol recommended for testing master seed virus.
Safety
Before proceeding with field trials, the safety of this new vaccine is tested in laboratory studies. The
following safety tests in particular should be carried out:
a)
The safety of the administration of one dose;
b)
The safety of the administration of an overdose (at least two doses of inactivated vaccine);
c)
The safety of the repeated administration of one dose.
The test is carried out for each approved route of administration. Use at least 10 adults (>4 months of
age) that are RHDV antibody free. Observe these animals for 21 days by evaluating the following life
parameters: general conditions and reactions, sensory condition, water and food consumption,
characteristics of faeces, and local abnormal reactions at the inoculum point. Record the body
temperature the day before vaccination, at vaccination, 4 hours after vaccination and then daily for
4 days; note the maximum temperature increase for each animal. No abnormal local or systemic
reaction should occur; the average body temperature increase should not exceed 1°C and no animal
should have a temperature rise greater than 2°C. A local reaction lasting less than 21 days may occur.
If the vaccine is intended for use in pregnant rabbits, administer the vaccine to not less than
10 pregnant does according to the schedule to be recommended. Prolong the observation period until
1 day after parturition. The does should remain in good health and there should not be abnormal local
or systemic reactions. No adverse effects on the pregnancy or on the offspring should be noted.
Batch potency
Use susceptible adult rabbits (>4 months old), free from antibodies against RHDV and reared in
suitable isolation conditions to ensure absence of contact with RHDV. Ten rabbits are vaccinated with
one full dose of vaccine given by the recommended route. Two other groups of five animals each are
vaccinated with 1/4 and 1/16 of the full dose, respectively. A fourth group of 10 unvaccinated rabbits is
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maintained as controls. All animals are challenged not less than 21 days post-vaccination by
intramuscular inoculation of a dose of RHDV containing at least 100 LD50 or presenting a HA titre
higher than 1/2560. Observe the rabbits for a further 21 days. The test is not valid if: a) during the
period between vaccination and challenge more than 10% of the vaccinated or more than 20% of
control rabbits show abnormal clinical signs or die from causes not attributable to the vaccine; b) less
than 70% of control rabbits died with typical signs of RHD. The vaccine complies with the test if: a) not
less than 90% of vaccinated rabbits show no signs of RHD; b) the mean antibody level of vaccinated
animals, is not significantly less than the level recorded in the protection test performed using as
vaccine the inactivated seed virus.
c)
Requirements for authorisation
The tests for safety, potency and sterility of the final product must be performed after bottling and packaging.
Thus, it is important that these two last manufacturing steps be performed following standardised good
manufacturing procedures. The tests are conducted by removing samples from a statistically determined
number of randomly taken multi-dose containers (20, 50 or 100 doses) of vaccine.
i)
Safety requirements
Target and non-target animal safety
Rabbit is the sole species susceptible to RHDV and in the interest of animal welfare, tests and trials
must be held only on target animals. The safety requirements of the final product for rabbits should be
verified in field studies on both fattening and breeder rabbits. At least 30 breeder rabbits, >4 months of
age, and 70 rabbits 30–45 days of age should be used. Breeder rabbits are vaccinated subcutaneously
at the back of the neck twice (at an interval of 3 weeks) with one dose. Fattening rabbit are vaccinated
either at 30 or 45 days of age. Animals are observed for 4 months from the first vaccination.
Unvaccinated animals are kept as controls.
The control of the safety of the vaccine in breeder rabbits is done by evaluating their reproductive
performance. The following parameters are considered: local or general reactions; total number of born
rabbits and the number of live rabbits born; percentage of mortality at the weaned period; average
weight of bunnies at the weaned period; daily consumption of food. The control of the safety of the
vaccine in fattened rabbits is done by evaluating their daily health. The following parameters are
considered: local or general reactions; individual weight increase from weaning (30 days) and every
15 days; daily consumption of feed; conversion index; mortality during the fattening period. Vaccinated
rabbits should not show any changes in their general conditions or abnormal local or systemic
reactions for the whole test duration.
The vaccine should not contain any ingredients that are likely to pose a risk for consumers of
vaccinated rabbits. However, as the inactivated vaccine contains a mineral oil adjuvant, there is an
associated risk that might arise from accidental self injection. Accidental injection can cause intense
swelling and severe consequences if expert medical advice is not sought promptly.
Reversion-to-virulence for attenuated/live vaccines
Reversion-to-virulence does not occur because it is an inactivated vaccine.
Environmental consideration
During the safety and efficacy field trials, interactions with other vaccines (e.g. vaccine against
myxomatosis) or pharmaceutical products (medicated feeding-stuffs containing antibiotics against
respiratory diseases and bacterial enteritis) should be checked and recorded. No interactions have
been reported to date.
The inactivated vaccine does not spread in the environment and, in previous trials, there were no signs
of ecotoxicity problems for the viral antigens. The risk of ecotoxicity caused by the use of vaccine is
zero because of the nature of the vaccine (inactivated vaccine for parenteral use). The vaccine
contains no ingredients likely to pose a risk to the environment. In addition, the vaccine is administered
by injection so environmental contamination is unlikely. To achieve the highest standard of safety in
accordance with good hygiene rules, the bottles must be dipped in an antiseptic solution after use.
ii)
Efficacy requirements
The efficacy should be tested in the laboratory with both challenge and serology tests. Forty rabbits
(20 vaccinated and 20 unvaccinated), at least 4 months of age, are challenged with virulent virus: all
the vaccinated animals must be protected, giving positive serological titres and all the control
unvaccinated animals must have died within the observation period.
The in-field efficacy of the vaccine may be determined by evaluating the seroconversion in blood
samples taken from both fattening and breeder rabbits at different check-points from vaccination. Titres
are measured by C-ELISA and anti-isotype IgM, IgA and IgG ELISAs.
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Before the first vaccination, the C-ELISA should confirm, in all rabbits, the total absence of anti-RHDV
antibodies or the presence of titres at the lower acceptance limit of the protection titre ≤1/10.
Vaccinated animals develop an RHDV protective immunity in a short period of time: in the serum of
infected animals, circulating antibodies are present just 3–4 days post-infection (IgM and IgA), whereas
in rabbits vaccinated with the inactivated and adjuvated vaccine, the first antibodies usually appear
after 7–10 days (only IgM). IgG appear after approximately 15–20 days. After vaccination there is very
low or no IgA production. As it is produced only during infection with the live virus following oro-nasal
dissemination, IgA could be considered to be a marker of contact with the field virus. The mucosal
immuno-system may also be involved in protection to the disease even if the vaccine is parentally and
not orally administered. This is suggested by oral challenge experiments in vaccinated rabbits when
IgA but no IgM appear very quickly in the serum. This suggests that B memory cells able to produce
IgA are already present at the mucosal level, which is usually the first site of replication of RHDV.
There is a definite correlation between the titre in the C-ELISA and the state of protection from the
disease, i.e. rabbits with titres higher than 1/10 for antibodies specifically induced by RHDV did not
show any sign of disease when challenged with virulent RHDV. In convalescent rabbits, serological
titres could be as high as 1/20480, whereas in vaccinated rabbits they are usually between 1/40 and
1/640 according to the time elapsed since vaccination. Maternal antibodies (IgG only) usually
disappear within 30 days of age in young rabbits born to vaccinated healthy does, but they last longer
(until 45–55 days of age) when rabbits are born to convalescent does, as the passive titres of young
are directly related to that of their mothers. This is true of young rabbits from industrial farms that are
weaned quite early (25–35 days of age), whereas in young wild rabbits, maternal antibodies can last
for 80 days (Forrester et al., 2002). In young rabbits (<35–40 days old), a low level of antibody (1/80–
1/320) could also be induced by an active infection not leading to disease, as commonly occurs in
animals of this age.
The data reported in the literature indicate the long-term duration of immunity induced by a single
vaccination (up to 15 months). At 9–12 months post-vaccination, titres are 2–4 times lower than
observed 2–3 weeks after vaccination. The booster effect, in the case of natural infection or revaccination, depends on the time elapsed since vaccination, i.e. it is lower 5–7 months post-vaccination
and higher in animals vaccinated before that time.
To exactly determine the duration and efficacy of immunity, it is advisable to carry out the following
test: 20 rabbits vaccinated once are divided into four groups and are serologically tested at monthly
intervals over a period of 1 year. Each group is inoculated with virulent RHDV at 3, 6, 9 months or
1 year post-vaccination. Challenge infection should produce increasing seroconversion, which is
directly related to the time that has elapsed since vaccination. The absence of clinical signs of disease
and mortality supports the efficacy of the vaccine.
iii)
Stability
Evidence should be provided to show that the vaccine passes the batch potency test at 3 months
beyond the suggested shelf life.
A suitable preservative is normally required for vaccine in multi-dose containers. Its persistence
throughout shelf life should be checked.
3.
Vaccines based on biotechnology
Several studies have been carried out on the expression of RHDV capsid protein in Escherichia coli, in vaccinia
virus, and in attenuated Myxoma virus (MV). Moreover, it has been shown by various authors that a recombinant
capsid protein, VP60, expressed in the baculovirus/Sf9 cell expression system, self assembled into VLPs that are
structurally and antigenically identical to RHD virions. While the fusion protein expressed in E. coli is highly
insoluble and of low immunogenicity, active immunisation can be achieved with VLPs obtained in the baculovirus
system or by using recombinant vaccinia, MV and canarypox, administered either intramuscularly or orally. In
particular, rabbits vaccinated with recombinant MV expressing the RHDV capsid protein were protected against
lethal RHDV and MV challenges. The resulting recombinant virus was also capable of spreading horizontally and
promoting protection of contact animals, thus providing the opportunity to immunise a wild rabbit population.
Similarly, the immunogenicity of VLPs administered by the oral route as an alternative to parenteral immunisation
offers an economical and practical way to administer a vaccine for mass immunisation of wild animals.
The VP60 structural protein has been also expressed in transgenic plants, either with a new plum pox virus
(PPV)-based vector (PPV-NK), or in transgenic potato plants under the control of a cauliflower mosaic virus 35S
promoter or a modified 35S promoter. In both cases the immunisation of rabbits with extracts of Nicotiana
clevelandii plants infected with the PPV-NK VP60 chimera and with leaf extracts from potatoes carrying this
modified 35S promoter, respectively, induced an efficient immune response that protected animals against a
lethal challenge with RHDV. However, at the present time, recombinant vaccines are not yet registered and
commercially available.
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In France, a vaccine has been commercialised that is a combination of a traditional inactivated liver-derived RHD
vaccine and a live attenuated Myxoma virus vaccine, and which can be administered by the intradermal route.
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MOSS S.R., TURNER S.L., TROUT R.C., WHITE P.J., HUDSON P.J., DESAI A, ARMESTO M, FORRESTER N.L. & GOULD E.A
(2002). Molecular epidemiology of Rabbit haemorrhagic disease virus. J. Gen. Virol., 83, 2461–2467.
NAGESHA H,S., MCCOLL K.A., COLLINS B.J., MORRISSY C.J., WANG L.F. & WESTBURY (2000). The presence of crossreactive antibodies to RHDV in Australian wild rabbits prior to the escape of the virus from quarantine. Arch. Virol.,
145, 749–757.
OHLINGER R.F., HAAS B., MEYERS G., WEILAND F. & THIEL H.J (1990). Identification and characterization of the virus
causing rabbit haemorrhagic disease. J. Virol., 64, 3331–3336.
ROBINSON A.J., KIRKLAND P.D., FORRESTER R.I., CAPUCCI L. & COOKE B.D. (2002). Serological evidence for the
presence of a calicivirus in Australian wild rabbits, Oryctolagus cuniculis, before the introduction of RHDV: its
potential influence on the specificity of a competitive ELISA for RHDV. Wildl. Res., 29, 655–662.
SCHIRRMEIER H., REIMANN I., KOLLNER B. & GRANZOW H. (1999). Pathogenic, antigenic and molecular properties of
rabbit haemorrhagic disease virus (RHDV) isolated from vaccinated rabbits: detection and characterization of
antigenic variants. Arch. Virol., 144, 719–735.
STOERCKLE-BERGER N., KELLER-BERGER B., ACKERMANN M. & EHRENSPERGER F. (1992). Immunohistological
diagnosis of rabbit haemorrhagic disease (RHD). J Vet. Med. [B], 39, 237–245.
STRIVE T., WRIGHT J.D. & ROBINSON A.J. (2009). Identification and partial characterisation of a new Lagovirus in
Australian wild rabbits. Virology, 384, 97–105.
WHITE P.J., TROUT R.C., MOSS S.R., DESAI A., ARMESTO M., FORRESTER N.L., GOULD E.A. & HUDSON P.J. (2004).
Epidemiology of rabbit haemorrhagic disease virus in the United Kingdom: evidence for seasonal transmission by
both virulent and avirulent modes of infection. Epidemiol. Infect., 132, 555–567.
WIRBLICH C., MEYERS G., OHLINGER V.F., CAPUCCI L., ESKENS U., HAAS B. & H.-J. THIEL (1994). European brown hare
syndrome virus: relationship to rabbit hemorrhagic disease virus and other caliciviruses. J. Virol., 68, 5164–5173.
*
* *
NB: There is an OIE Reference Laboratory for Rabbit haemorrhagic disease
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for rabbit haemorrhagic disease
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
SECTION 2.7.
OVIDAE AND CAPRINAE
CHAPTER 2.7.1.
BORDER DISEASE
SUMMARY
Border disease (BD) is a viral disease of sheep and goats first reported in sheep in 1959 from the
border region of England and Wales, and since recorded world-wide. Distribution of the virus is
world-wide. Prevalence rates vary in sheep from 5% to 50% between countries and from region to
region within countries. Clinical signs include barren ewes, abortions, stillbirths and the birth of
small weak lambs. Affected lambs can show tremor, abnormal body conformation and hairy fleeces
(so-called ‘hairy-shaker’ or ‘fuzzy’ lambs) and the disease has been referred to as ‘hairy shaker
disease’. Vertical transmission plays an important role in the epidemiology of the disease. Infection
of fetuses can result in the birth of persistently infected (PI) lambs. These PI lambs are viraemic,
antibody negative and constantly excrete virus. The virus spreads from sheep to sheep, with PI
animals being the most potent source of infection. Infection in goats is less common with abortion
being the main presenting sign. In many regions the commonest cause of BD is the pestivirus
border disease virus (BDV), but in some parts of the world, bovine viral diarrhoea virus (BVDV) may
be a more common cause of BD. The source of BVDV for sheep often is close contact with cattle.
It is important to identify the viraemic PI animals so that they will not be used for breeding or trading
purposes. Serological testing is insufficient. It is generally considered that serologically positive,
nonviraemic sheep are ‘safe’, as latent infections are not known to occur in recovered animals.
Identification of the agent: BD virus (BDV) is a Pestivirus in the family Flaviviridae and is closely
related to classical swine fever virus and BVDV. Nearly all isolates of BDV are noncytopathogenic
in cell culture. There are no defined serotypes but virus isolates exhibit considerable diversity.
Three distinct antigenic groups, plus two further separate genotypes, have been identified.
Apparently healthy PI sheep resulting from congenital infection can be identified by isolation and
immunostaining of noncytopathogenic virus from blood or sera in laboratory cell cultures. Rapid
direct methods to identify PI sheep include detection of viral antigen or viral RNA in leukocytes and
immunohistochemical demonstration of viral antigen in skin biopsies. The demonstration of virus is
less reliable in lambs younger than 2 months that have received colostral antibody. Acute infection
is usually subclinical and viraemia is transient and difficult to detect. From dead animals, the
isolation of virus from tissues of aborted or stillborn lambs is difficult, but tissues from PI sheep
contain high levels of virus, which can be easily detected by isolation and direct methods.
Serological tests: Acute infection with BDV is best confirmed by demonstrating seroconversion
using paired or sequential samples from several animals in the group. The enzyme-linked
immunosorbent assay and virus neutralisation test are the most commonly used antibody detection
methods.
Requirements for vaccines and diagnostic biologicals: There is no standard vaccine for BDV,
but a commercial killed whole-virus vaccine has been produced. Ideally, such a vaccine should be
suitable for administration to females before breeding for prevention of transplacental infection. The
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use of BVDV vaccines has been advocated, but the antigenic diversity of BD viruses must be
considered.
BD viruses have contaminated several modified live veterinary vaccines produced in sheep cells or
containing sheep serum. This potential hazard should be recognised by manufacturers of biological
products.
A. INTRODUCTION
Border disease virus (BDV) is a Pestivirus of the family Flaviviridae and is closely related to classical swine fever
virus (CSFV) and bovine viral diarrhoea virus (BVDV). Pestivirus taxonomy is quite problematic at present. There
are four recognised species, namely – CSFV, BVDV types 1 and 2 and BDV (Theil et al., 2005). While CSF
viruses are predominantly restricted to pigs, examples of the other three species have all been recovered from
sheep, with the majority of isolates being BD viruses (Vilcek et al., 1997). Nearly all virus isolates of BDV are
noncytopathogenic, although occasional cytopathic viruses have been isolated (Vantsis et al., 1976). BDV
spreads naturally among sheep by the oro-nasal route and by vertical transmission. It is principally a cause of
congenital disease in sheep and goats, but can also cause acute and persistent infections. Infection is less
common in goats, in which persistent infection is rare as abortion is the main presenting sign. Sheep can become
infected with BVDV from cattle (Carlsson, 1991), and in some countries, BVDV can be a more common cause of
BD than BDV. Pigs may also be infected by pestiviruses other than CSFV and antibodies to BDV in pigs may
interfere with tests for the diagnosis of CSF (Oguzoglu et al., 2001). Several new BD viruses from sheep, goats
and Pyrenean chamois (Rupicapra pyrenaica pyrenaica) have been described recently. Phylogenetic analysis
using computer-assisted nucleotide sequence analysis suggests that genetic variability among BD viruses is
greater than within each of the other pestivirus species. Four distinguishable genogroups of BDV have been
described as well as putative novel pestivirus genotypes from Tunisian sheep and a goat (Becher et al., 2003;
Vilcek & Nettleton, 2006). The chamois BD virus is similar to isolates from sheep in the Iberian Peninsula
(Valdazo-Gonzalez et al., 2007). This chapter describes BDV infection in sheep.
a)
Acute infections
Healthy newborn and adult sheep exposed to BDV experience only mild or inapparent disease. Slight fever
and a mild leukopenia are associated with a short-lived viraemia detectable between days 4 and 11 postinfection, after which virus neutralising antibody appears in the serum (Thabti et al., 2002).
Acute infections are best diagnosed serologically using paired sera from a representative number of sheep.
Occasional BDV isolates have been shown to produce high fever, profound and prolonged leukopenia,
anorexia, conjunctivitis, nasal discharge, dyspnoea and diarrhoea, and 50% mortality in young lambs. One
such isolate was recovered from a severe epidemic of BD among dairy sheep in 1984 (Chappuis et al.,
1986). A second such isolate was a BDV contaminant of a live CSFV vaccine (Wensvoort & Terpstra, 1988).
b)
Fetal infection
The main clinical signs of BD are seen following the infection of pregnant ewes. While the initial maternal
infection is subclinical or mild, the consequences for the fetus are serious. Fetal death may occur at any
stage of pregnancy, but is more common in fetuses infected early in gestation. Small dead fetuses may be
resorbed or their abortion may pass unnoticed as the ewes continue to feed well and show no sign of
discomfort. As lambing time approaches, the abortion of larger fetuses, stillbirths and the premature births of
small, weak lambs will be seen. Confirmation that an abortion or stillbirth is due to BDV is often difficult to
establish, but virus may be isolated from fetal tissues in some cases. In aborted fetuses, it is also possible to
detect virus by immunohistochemistry of brain, thyroid and other tissues (Thur et al., 1997). Samples of fetal
fluids or serum should be tested for BDV antibody.
During lambing, an excessive number of barren ewes will become apparent, but it is the diseased live lambs
that present the main clinical features characteristic of BD. The clinical signs exhibited by BD lambs are very
variable and depend on the breed of sheep, the virulence of the virus and the time at which infection was
introduced into the flock. Affected lambs are usually small and weak, many being unable to stand. Nervous
signs and fleece changes are often apparent. The nervous signs of BD are its most characteristic feature.
The tremor can vary from violent rhythmic contractions of the muscles of the hindlegs and back, to barely
detectable fine trembling of the head, ears, and tail. Fleece abnormalities are most obvious in smoothcoated breeds, which develop hairy fleeces, especially on the neck and back. Abnormal brown or black
pigmentation of the fleece may also be seen in BD-affected lambs. Blood samples to be tested for the
presence of BDV and/or antibody should be collected into anticoagulant from suspect lambs before they
have received colostrum. Once lambs have ingested colostrum, it is difficult to detect virus until they are
2 months old and maternal antibody levels have waned. However, during this period, it may be possible to
detect viral antigen in skin biopsies, by immunohistochemistry, in washed leukocytes by enzyme-linked
immunosorbent assay (ELISA) or by reverse-transcription polymerase chain reaction (RT-PCR).
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With careful nursing, a proportion of BD lambs can be reared, although deaths may occur at any age. The
nervous signs gradually decline and can disappear by 3–6 months of age. Weakness, and swaying of the
hind-quarters, together with fine trembling of the head, may reappear at times of stress. Affected lambs often
grow slowly and under normal field conditions many will die before or around weaning time. In cases where
losses at lambing time have been low and no lambs with obvious signs of BD have been born, this can be
the first presenting sign of disease.
Some fetal infections occurring around mid-gestation can result in lambs with severe nervous signs,
locomotor disturbances and abnormal skeletons. Such lambs have lesions of cerebellar hypoplasia and
dysplasia, hydranencephaly and porencephaly resulting from necrotising inflammation. The severe
destructive lesions appear to be immune mediated, and lambs with such lesions frequently have high titres
of serum antibody to BDV. Most lambs infected in late gestation are normal and healthy and are born free
from virus but with BDV antibody. Some such lambs can be weak and may die in early life (Barlow &
Patterson, 1982).
c)
Persistent viraemia
When fetuses survive an infection that occurs before the onset of immune competence, they are born with a
persistent viraemia. The ovine fetus can first respond to an antigenic stimulus between approximately
60 and 85 days of its 150-day gestation period. In fetuses infected before the onset of immune competence,
viral replication is uncontrolled and 50% fetal death is common. In lambs surviving infection in early
gestation, virus is widespread in all organs. Such lambs appear to be tolerant of the virus and have a
persistent infection, usually for life. A precolostral blood sample will be virus positive and antibody negative.
Typically, there is no inflammatory reaction and the most characteristic pathological changes are in the
central nervous system (CNS) and skin. Throughout the CNS, there is a deficiency of myelin, and this
causes the nervous signs. In the skin, primary wool follicles increase in size and the number of secondary
wool follicles decreases, causing the hairy or coarse fleece.
Persistently viraemic sheep can be diagnosed by virus isolation/detection in a blood sample. Viraemia is
readily detectable at any time except within the first 2 months of life, when virus is masked by colostral
antibody; however, the virus may be detected in washed leukocytes during this period, and in animals older
than 4 years, some of which develop low levels of anti-BDV antibody (Nettleton et al., 1992). Although virus
detection in blood during an acute infection is difficult, persistent viraemia should be confirmed by retesting
animals after an interval of at least 3 weeks.
Some viraemic sheep survive to sexual maturity and are used for breeding. Lambs born to these infected
dams are always persistently viraemic. Persistently viraemic sheep are a continual source of infectious virus
to other animals and their identification is a major factor in any control programme. Sheep being traded
should be screened for the absence of BDV viraemia.
Usually persistently infected (PI) rams have poor quality, highly infective semen and reduced fertility. All
rams used for breeding should be screened for persistent BDV infection on a blood sample. Semen samples
can also be screened for virus, but virus isolation is much less satisfactory than from blood due to the toxicity
of semen for cell cultures. RT-PCR for detecting pestivirus nucleic acid may be justifiable on semen from
some rams.
d)
Late-onset disease in persistently viraemic sheep
Some PI sheep housed apart from other animals spontaneously develop intractable diarrhoea, wasting,
excessive ocular and nasal discharges, sometimes with respiratory distress. At necropsy such sheep have
gross thickening of the distal ileum, caecum and colon resulting from focal hyperplastic enteropathy.
Cytopathic BDV can be recovered from the gut of these lambs. With no obvious outside source of cytopathic
virus, it is most likely that such virus originates from the lamb's own virus pool. Other PI sheep in the group
do not develop the disease. This syndrome, which has been produced experimentally and recognised in
occasional field outbreaks of BD, has several similarities with bovine mucosal disease (Nettleton et al.,
1992).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent (the prescribed test for international trade)
There is no designated OIE reference laboratory for BDV, but the reference laboratories for BVDV or CSFV will
be able to provide advice (see Table given in Part 4 of this Terrestrial Manual). One of the most sensitive proven
methods for identifying BDV remains virus isolation. Direct immunofluorescence or other immunohistochemical
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techniques on frozen tissue sections as well as antigen-detecting ELISA and conventional and real-time RT-PCR
are also valuable methods for identifying BDV-infected animals.
a)
Virus isolation
It is essential that laboratories undertaking virus isolation have a guaranteed supply of pestivirus-free
susceptible cells and fetal bovine serum (FBS), or equivalent, that contain no anti-pestivirus activity and no
contaminating virus. It is important that a laboratory quality assurance programme be in place.
The virus can be isolated in a number of primary or secondary ovine cell cultures (e.g. kidney, testes, lung).
Ovine cell lines for BDV growth are rare. Semicontinuous cell lines derived from fetal lamb muscle (FLM),
whole embryo (Thabti et al., 2002) or sheep choroid plexus can be useful, but different lines vary
considerably in their susceptibility to the virus. Ovine cells have been used successfully for the isolation and
growth of BD viruses and BVDV types 1 and 2 from sheep. In regions where sheep may become infected
with BVD viruses from cattle, a virus isolation system using both ovine and bovine cells could be optimal.
Several bovine cell cultures may be suggested, including testicular, embryonic tracheal or turbinate cells, or
a susceptible continuous kidney cell line. However, bovine cells are insensitive for the primary isolation and
growth of some BD viruses, so reliance on bovine cells alone is inadvisable.
From live animals, serum can be tested for the presence of infectious virus, but the most sensitive way to
confirm pestivirus viraemia is to wash leukocytes repeatedly (at least three times) in culture medium before
co-cultivating them with susceptible cells for 5–7 days. Cells are frozen and thawed once and an aliquot
passaged onto further susceptible cells grown on cover-slips, chamber slides or plastic plates. The cells are
stained, 3–4days later, for the presence of pestivirus using an immunofluorescence or immunoperoxidase
test. Tissues should be collected from dead animals in virus transport medium (10% [w/v]). In the laboratory,
the tissues are ground, centrifuged to remove debris, and the supernatant passed through 0.45 µm filters.
Spleen, thyroid, thymus, kidney, brain, lymph nodes and gut lesions are the best organs for virus isolation.
Semen can be examined for the presence of BDV, but raw semen is strongly cytotoxic and must be diluted,
usually at least 1/10 in culture medium. As the major threat of BDV-infected semen is from PI rams, blood is
a more reliable clinical sample than semen for identifying such animals. There are many variations in virus
isolation procedures. All should be optimised for maximum sensitivity using a standard reference virus
preparation and, whenever possible, recent BDV field isolates. A practical sensitive isolation procedure is
outlined below:
i)
Cultures with subconfluent or newly confluent monolayers of susceptible ovine cells are washed at
least twice with Hanks balanced salt solution to remove growth medium before being inoculated with
approximately 0.2 ml of sample, which is allowed to adsorb for 2 hours at 37°C.
ii)
Cultures are washed with at least 2 ml medium. This is discarded and an appropriate volume of culture
maintenance medium is added.
iii)
Cultures are incubated for 5–7 days at 37°C. They are examined microscopically on a daily basis and
evidence of cytopathic effect (CPE) is recorded.
iv)
Cultures are frozen at –70°C and then thawed for passage, as before, on to fresh cultures.
v)
3–4 days later, cells growing on glass are fixed in cold acetone for 15 minutes while cells growing on
plastic are fixed as described in the virus neutralisation test section below. Fixed cells are stained using
an indirect or direct immunofluorescence method. Essential controls must include known negative cells
and cells growing standard cytopathic and noncytopathic BDV strains.
vi)
The cells are examined under a UV microscope for the diffuse cytoplasmic fluorescence that is
characteristic of pestiviruses.
Immunoperoxidase staining can also be used on cover-slips, chamber slides as well as microtitre plates (see
method under virus neutralisation [VN] test below). Frozen and thawed cultures can also be tested in an
antigen detection ELISA system that employs monoclonal antibodies (MAbs) against epitopes on the
conserved nonstructural NS 2-3 protein. Staining for noncytopathic pestiviruses will usually detect virus at
the end of the first passage, but in order to detect slow-growing viruses in poorly permissive cells two
passages are desirable.
b)
Immunohistochemistry
Viral antigen demonstration is possible in most of the tissues of PI animals (Braun et al., 2002; Thur et al.,
1997). This should be done on acetone-fixed frozen tissue sections (cryostat sections) or paraffin wax
embedded samples using appropriate antibodies. Panpestivirus-specific antibodies with NS2-3 specificity
are suitable. Tissues with a high amount of viral antigen are brain, thyroid gland and oral mucosa. Skin
biopsies have been shown to be useful for in-vivo diagnosis of persistent BDV infection.
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Chapter 2.7.1. — Border disease
c)
Enzyme-linked immunosorbent assay for antigen detection
The first ELISA for pestivirus antigen detection was described for detecting viraemic sheep. This has now
been modified into a double MAb capture ELISA for use in sheep and cattle. Two capture MAbs are bound
to wells in microtitre plates, and two other MAbs, conjugated to peroxidase, serve as detector MAbs
(Entrican et al., 1994). The test is most commonly employed to identify PI viraemic sheep using washed,
detergent-lysed blood leukocytes. The sensitivity is close to that of virus isolation and it is a practical method
for screening high numbers of blood samples. As with virus isolation, high levels of colostral antibody can
mask persistent viraemia. The ELISA is more effective than virus isolation in the presence of antibody, but
may give false-negative results in viraemic lambs younger than 2 months old. The ELISA is usually not
sensitive enough to detect acute BDV infections on blood samples. As well as for testing leukocytes, the
antigen ELISA can also be used on tissue suspensions, especially spleen, from suspected PI sheep and, as
an alternative to immunofluorescence and immunoperoxidase methods, on cell cultures. Several pestivirus
ELISA methods have been published and commercial kits are now available for detecting BDV. ELISAs
employing MAbs recognising epitopes on the conserved non-structural NS2-3 should recognise all strains of
BDV. ELISAs relying on MAbs recognising epitopes on structural proteins such as Erns, that are used for
BVDV detection in cattle, are unsuitable for the diagnosis of BDV viraemia in sheep.
d)
Nucleic acid detection methods
The complete genomic sequences of three BD viruses have been determined and compared with those of
other pestiviruses (Becher et al., 1998; Ridpath & Bolin, 1997). Phylogenetic analysis shows BD viruses to
be more closely related to CSFV than to BVDV (Becher et al., 2003; Van Rijn et al., 1997; Vilcek & Nettleton,
2006; Vilcek et al., 1997). RT-PCR for diagnosing pestivirus infection is now used widely. Various formats
are described. Basic RT-PCR protocols involve the following stages:
i)
Total RNA is isolated by phenol-chloroform, TRIZOL, guanidine isothiocyanate (GITC), or a
commercially available spin column or magnetic bead separation methods. NOTE: many of these
chemicals are highly toxic; adhere to the manufacturers’ safety procedures.
ii)
RT-PCR is performed. This is a two-stage reaction that consists of:
a)
Reverse transcription to produce single-stranded cDNA from viral RNA;
b)
Subsequent PCR amplification of the cDNA to produce readily detectable amounts of doublestranded DNA.
This process may be done as two separate reactions, each done in a separate PCR tube (two-step
RT-PCR), or as two stages in a single PCR tube (one-step RT-PCR). In a two-step format, either
random hexamers or specific primers may be used to prime the RT stage; in the one-step format only
specific primers may be used.
iii)
Specific product is detected by one of the following methods:
a) Use of BD-specific primers in the RT-PCR, with visualisation by agarose gel electrophoresis,
ethidium bromide staining and UV transillumination to demonstrate the correct sized amplicon.
NOTE: ethidium bromide is highly toxic; adhere to manufacturer’s recommendations for handling.
NOTE: UV transillumination must be carried out taking appropriate precautions to minimise skin
exposure.
b) Nested PCR, using pan-pestivirus primers (usually directed to the 5’UTR region) in a primary PCR,
followed by specific BD virus primers in a secondary (nested) PCR. Such assays typically employ
approximately 25 cycles in the primary PCR and 30–35 cycles in the nested PCR. Amplicons are
detected by visualisation by agarose gel electrophoresis as above. These assays increase
specificity and sensitivity but are more susceptible to contamination.
c) Real-time RT-PCR, using specific BD primers and/or fluorophore-labelled oligonucleotide probe to
detect BD. This method has advantages in specificity and prevention of contamination. It is also
possible to carry out a nested form of real-time RT-PCR.
Oligonucleotide primer/probe design is critical to the success of these assays, due to the genetic variability
of BDV isolates. Panpestivirus primers are valuable for detecting and typing all species of Pestivirus
(Sandvik et al., 1997; Vilcek et al., 1994), and can be combined with sequencing if required for specificity or
epidemiological investigation. Specific primers for the specific recognition of BD viruses have also been
described (Fulton et al., 1999; Vilcek & Paton, 2000; Willoughby et al., 2006). Using a closed one-tube RTPCR with fluorescent probes reduces the potential for cross-contamination of diagnostic samples
(McGoldrick et al., 1999; Willoughby et al., 2006). The development of a real-time RT-PCR allows the rapid
simultaneous detection and typing of ovine pestiviruses (Willoughby et al., 2006). Important applications of
RT-PCR methods include the detection of viral RNA in fetal tissues and in cell culture constituents or
vaccines (Vilcek, 2001); it may also prove valuable for detecting virus when BDV-specific antibodies are
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present. Validation of the RT-PCR is in process. The precautions to be taken with RT-PCR have been
covered in Chapter 1.1.5 Principles and methods of validation of diagnostic assays for infectious diseases.
2.
Serological tests
Antibody to BDV is usually detected in sheep sera using VN or an ELISA. The less sensitive agar gel
immunodiffusion (AGID) test may also be used. Control positive and negative reference sera must be included in
every test. These should give results within predetermined limits for the test to be considered valid. Single sera
can be tested to determine the prevalence of BDV in a flock, region or country. For diagnosis, however, acute and
convalescent sera are the best samples for confirming acute BDV infection. Repeat sera from one animal should
always be tested alongside each other on the same plate.
a)
Virus neutralisation test
A standard cytopathic strain of BDV (e.g. Moredun strain) can be used for the VN test with semicontinuous
cells such as FLM. An outline protocol is given below.
i)
Test and control sera are heat-inactivated for 30 minutes at 56°C.
ii)
From a starting serum dilution of 1/4, serial twofold dilutions of the test sera are made in cell culture
growth media in a cell culture-grade flat-bottomed 96-well microtitre plate. For each sample two or four
wells are used at each dilution depending on the accuracy required. The range of dilution can also
vary. It is common to screen sera initially at a dilution of 1/4 and titrate out positive sera. To screen
sera a minimum of four wells is required. The standard working volume is 25 µl: 25 µl of the diluted
serum is added to each well; 25 µl of media is added to each of the lower two control wells and 25 µl of
media containing 100 TCID50 (50% tissue culture infective dose) of virus is added to each of the two
upper test wells. Control positive and negative sera and a virus titration are included in every test.
iii)
Plates are sealed with nontoxic plate sealers or lids, and incubated at 37°C for 1 hour.
iv)
100 µl of a cell suspension with a count of 2 × 105 cells/ml is added to every well. The FBS or
equivalent serum used for cell growth must be free from antibody to BDV.
v)
The plate is sealed or incubated in a humid chamber in 5% CO2 for 4 days at 37°C.
vi)
The wells are examined microscopically for CPE. In the control wells of the test sera, cell degeneration
will be due to toxicity. Further dilution of toxic sera can be attempted, but it may not be possible to
obtain reliable results with occasional sera. The VN titre for each serum is the dilution at which the
virus is neutralised in 50% of the wells. This can be calculated by the Spearman–Kärber method. A
seronegative animal will show no neutralisation at the lowest dilution (i.e. 1/4).
The choice of test virus is difficult due to antigenic diversity among pestiviruses (Dekker et al., 1995;
Nettleton et al., 1998) Standard strains of cytopathic BVD viruses and bovine cells can be used. Oregon
C24V results correlate better with Moredun BDV than results with the NADL strain. No single strain is ideal.
A local strain that gives the highest antibody titre with a range of positive sheep sera should be used. The
VN test can also be used with noncytopathic viruses when the following immunoperoxidase staining system
is used after step v above:
i)
The culture medium is removed and the cells are washed gently with warm phosphate buffered saline
(PBS), air-dried and cooled to 4°C.
ii)
The cells are fixed by quickly adding to all wells 95% acetone (in water) previously cooled to –20°C.
The plates are held at –20°C for 30 minutes and should not be stacked or allowed to warm as etching
of the plastic may occur.
iii)
The acetone is removed and the plates are dried quickly in a cool environment.
iv)
50 µl of BDV antiserum is added to all wells at a predetermined dilution in PBS with 1% Tween 80
(PBST). The plates are incubated at 37°C for 30 minutes in a humid atmosphere.
v)
The plates are emptied and washed three times with PBST.
vi)
The wells are drained and an appropriate anti-species serum conjugated to peroxidase at a
predetermined dilution is added, and the plates are left for 30 minutes at 37°C in a humid atmosphere.
vii)
The plates are emptied and washed three times with PBST.
viii) The plates are drained well and 50 µl of activated substrate, e.g. 3-amino-9-ethyl carbazole (AEC) is
added. AEC stock solution is: AEC (0.1 g) dissolved in dimethyl formamide (15 ml). For use add stock
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(0.3 ml) to membrane-filtered 0.05 M acetate buffer, pH 5.0, (4.7 ml), and then add 30% H2O2 (5 µl).
NOTE. This solution is toxic and should be handled with adequate precautions.
ix)
The plates are incubated at room temperature and known virus-positive control wells are monitored for
development of specific red-brown cytoplasmic staining. When staining is complete the substrate is
removed carefully and the wells are washed thoroughly with tap water. Leaving the tap water in the
wells, the plates are examined microscopically for virus-containing wells.
x)
The VN titre is calculated as above using the Spearman–Kärber method.
xi)
Alternatively, the test can be performed using direct fluorescein-isothiocyanate conjugate staining.
Occasionally there may be a need to determine whether antibody in a flock is against a virus belonging to a
particular Pestivirus serogroup. A differential VN test can be used in which sera are titrated out against
representative viruses from each of the four Pestivirus groups, i.e. BDV, BVDV types 1 and 2, and CSFV.
Maximum titre will identify the infecting serotype and the spectrum of cross-reactivity with the other
serotypes will also be revealed.
b)
Enzyme-linked immunosorbent assay
An MAb-capture ELISA for measuring BDV antibodies has been described. Two panpestivirus MAbs that
detect different epitopes on the immunodominant nonstructural protein NS 2/3 are used to capture
detergent-lysed cell-culture grown antigen. The results correlate qualitatively with the VN test (Fenton et al.,
1991).
Antigen is prepared as follows: Use eight 225 cm2 flasks of newly confluent FLM cells; four flasks will be
controls and four will be infected. Wash the flasks and infect four with a 0.01–0.1 m.o.i. (multiplicity of
infection) of Moredun cytopathic BDV. Allow the virus to adsorb for 2 hours at 37°C. Add maintenance media
containing 2% FBS (free from BDV antibody), and incubate cultures for 4–5 days until CPE is obvious. Pool
four control flask supernatants and separately pool four infected flask supernatants. Centrifuge at 3000 g for
15 minutes to pellet cells. Discard the supernatants. Retain the cell pellets. Wash the flasks with 50 ml of
PBS and repeat the centrifugation step as above. Pool all the control cell pellets in 8 ml PBS containing 1%
Nonidet P40 and return 2 ml to each control flask to lyse the remaining attached cells. Repeat for infected
cells. Keep the flasks at 4°C for at least 2 hours agitating the small volume of fluid on the cells vigorously
every 30 minutes to ensure total cell detachment. Centrifuge the control and infected antigen at 12,000 g for
5 minutes to remove the cell debris. Supernatant antigens are stored at –70°C in small aliquots.
•
Test procedure
i)
The two MAbs are diluted to a predetermined dilution in 0.05 M bicarbonate buffer, pH 9.6. All wells of a
suitable ELISA-grade microtitre plate (e.g. Nunc maxisorb, Greiner 129b) are coated overnight at 4°C.
ii)
After washing three times in PBST, a blocking solution of PBST containing 10% horse serum (PBSTH)
is added to all wells, which are incubated at 37°C for 1 hour.
iii)
The antigen is diluted to a predetermined dilution in PBSTH and alternate rows of wells are coated with
virus and control antigens for 1 hour at 37°C. The plates are then washed three times in PBST before
addition of test sera.
iv)
Test sera are diluted 1/50 in PBSTH and added to duplicate virus and duplicate control wells for 1 hour
at 37°C. The plates are then washed three times in PBST.
v)
Anti-ovine IgG peroxidase conjugate is diluted to a predetermined dilution in PBSTH and added to all
wells for 1 hour at 37°C. The plates are washed three times in PBST.
vi)
A suitable activated enzyme substrate, such as ortho-phenylene diamine (OPD) or tetramethyl blue
(TMB), is added noting the manufacturer’s toxicity warning. After colour development, the reaction is
stopped with sulphuric acid and the absorbance read on an ELISA plate reader. The mean value of the
two control wells is subtracted from the mean value of the two virus wells to give the corrected
absorbance for each serum. Results are expressed as corrected absorbance with reference to the
corrected absorbance of known positive and negative sera. Alternatively, ELISA titres can be
extrapolated from a standard curve of a dilution series of a known positive reference serum.
If antigens of sufficient potency can be produced the MAb capture stage can be omitted. In this case
alternate rows of wells are coated with virus and control antigen diluted to a predetermined dilution in
0.05 M bicarbonate buffer, pH 9.6, overnight at +4°C. The plates are washed and blocked as in step ii
above. After washing, diluted test sera are added and the test proceeds from step iv as above.
c)
Agar gel immunodiffusion test
The AGID test was first used to demonstrate an immunological relationship between BD, BVD and CSF
viruses.
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The Oregon C24V strain of BVDV grown on calf testis cells has been used to detect antibody in sheep.
Suitable antigen can be prepared using medium harvested from cells showing early CPE. Concentration of
the medium approximately 100-fold by dialysis against polyethylene glycol (PEG) is required. Alternatively,
PEG 6000 can be added to sonicated virus/cell suspensions at the rate of 8% (w/v). After constant stirring
overnight at 4°C, the precipitate is removed by centrifugation at 1800 g for 1 hour. The supernatant is
decanted thoroughly and the precipitate resuspended to 1% of the original virus/cell culture volume in
distilled water. The resuspended precipitate is centrifuged at 286,000 g for 2 hours and the supernatant
withdrawn for use as antigen. The precipitate is discarded.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
To be useful, a BDV vaccine should be effective when administered to female sheep before breeding to prevent
transplacental infection. Experimental and commercial inactivated whole virus BDV vaccines have been produced
in Europe (Brun et al., 1993; Vantsis et al., 1980).
Pestivirus contaminants of modified live virus vaccines have been found to be a cause of serious disease
following their use in pigs, cattle, sheep and goats. Contaminated vaccines have included those used for the
control of Aujesky’s disease, CSF, rotavirus, coronavirus, rinderpest, sheep pox and contagious pustular
dermatitis. The insidious ability of pestiviruses to cross the placenta, and thus establish PI animals, gives them
the potential to contaminate vaccines through cells, serum used as medium supplement, or seed stock virus. As
nearly all isolates of pestiviruses are noncytopathic, they will remain undetected unless specific tests are carried
out.
1.
Seed management
a)
Characterisation of the seed
An ideal vaccine should contain a strain or strains of virus that give protection against all sheep pestiviruses.
Recent evidence is that three antigenically distinguishable groups of pestiviruses infect sheep. One group is
represented by the Moredun reference strain of BDV; the second group contains viruses similar to the
majority of cattle BVDV strains (BVDV type 1); and the third group contains the less common BVDV (type 2)
strains (Wensvoort et al., 1989). More recently, ovine pestivirus isolates have been divided on the basis of
phylogenetic and antigenic analysis into BDV-1, BDV-2 and BDV-3 genotypes (Becher et al., 2003).
Phylogenetic analysis alone suggests that a BDV Italian caprine isolate and the chamois/Iberian sheep
isolates represent two further genotypes (Vilcek & Nettleton, 2006). Further cross-neutralisation studies are
required to determine the significance of these findings. Nevertheless it would appear that any BDV vaccine
should contain at least a representative of the BDV and BVDV (type 1) groups. Characterisation of the
biologically cloned vaccine viruses should include typing with MAbs and genotyping (Paton et al., 1995).
b)
Culture
A variety of ruminant cell cultures can be used. Optimal yields depend on the cell type and isolate used. A
commercial BDV vaccine containing two strains of virus is prepared on ovine cell lines (Brun et al., 1993).
Cells must be produced according to a seed-lot system from a master cell seed (MCS) that has been shown
to be free from all contaminating microorganisms. Vaccine should only be produced in cells fewer than
20 passages from the MCS. Control cells from every passage should be checked for pestivirus
contamination.
c)
Validation as a vaccine
All vaccines should pass standard tests for safety and efficacy. Safety testing of inactivated BDV vaccines
should include monitoring of all vaccine components for contaminating pestiviruses.
Efficacy tests of BDV vaccines should demonstrate their ability to prevent transplacental spread of virus.
Effective challenge of vaccinated pregnant ewes at 50–60 days gestation has been achieved by intranasal
installation of virus or by mixing with PI sheep (Brun et al., 1993).
2.
Method of manufacture
Inactivated vaccines have been prepared using conventional laboratory techniques with stationary or rolled cell
cultures. Inactivants have included formalin and beta-propriolactone. Adjuvants have included aluminium
hydroxide and oil (Brun et al., 1993; Vantsis et al., 1980).
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3.
In-process control
Cultures should be inspected daily to ensure they are free from gross bacterial contamination and that any CPE
observed is appropriate to the cytopathic virus being grown. No CPE should be observed in cultures being used
to grow noncytopathic strains of virus.
4.
Batch control
a)
Sterility
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
b)
Safety
Samples from inactivated vaccines should be tested rigorously for viable virus. Samples of the product
should be passaged several times in sensitive cell cultures to ensure absence of live BDV. This in-vitro
monitoring can be augmented by injecting two BDV-seronegative sheep with 20 doses of unformulated
antigen as part of a standard safety test. Presence of live virus will result in the development of a more
convincing serological response than will occur with inactivated virus alone. The sheep sera can also be
examined for antibody to other prescribed agents.
c)
Potency
Vaccine potency is also best tested in seronegative sheep in which the development and level of antibody is
measured. An indirect measure of potency is given by the level of virus infectivity prior to inactivation. The
antigen content following inactivation can be assayed by MAb-capture ELISA and related to the results of
established in-vivo potency results. As recommended for potency testing of BVDV vaccine in cattle it should
be demonstrated that the vaccine can prevent transplacental transmission of BDV in pregnant sheep.
d)
Duration of immunity
No information is available on duration of immunity following vaccination. Inactivated vaccines are unlikely to
provide sustained levels of immunity and it is likely that after an initial course of two or three injections
annual booster doses may be required. Insufficient information is available to determine any correlation
between vaccinal antibody titres in the dam and fetal protection.
e)
Stability
There is little information on the stability of BDV vaccines. Inactivated vaccines could be expected to have at
least a 1 year shelf life when protected from light and stored at 4°C.
f)
Preservatives
Preservatives may be added to multidose vaccine containers subject to the approval of the Control Authority.
g)
Precautions (hazards)
BDV is not considered to be a hazard to human health. Standard good microbiological practice should be
used when handling the virus.
5.
Tests on the final product
a)
Safety
In-vitro test only.
b)
Potency
In-vitro antigen content test.
REFERENCES
BARLOW R.M. & PATTERSON D.S.P. (1982). Border disease of sheep: a virus-induced teratogenic disorder. Adv.
Vet. Med. (Suppl. J. Vet. Med.), 36, 1–87.
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BECHER P., AVALOS-RAMIREZ R., ORLICH M., CEDILLO ROSALES S., KONIG, M., SCHWEIZER M., STALDER H., SCHIRRMEIR
H & THIEL H.-J. (2003). Genetic and antigenic characterisation of novel pestivirus genotypes; Implications for
classification. Virology, 311, 96–104.
BECHER P., ORLICH M. & THIEL H.-J. (1998). Complete genomic sequence of border disease virus a pestivirus from
sheep. J. Virol., 72, 5165–5173.
BRAUN U., HILBE M., EHRENSPERGER F., SALIS F., ALTHER P., STRASSER M., STALDER H.P. & PETERHANS E. (2002).
Border Disease in einem Schafbetrieb. Schweiz. Arch. Tierheilk., 144, 419–426.
BRUN A., LACOSTE F., REYNAUD G., KATO F. & SAINT-MARC B. (1993). Evaluation of the potency of an inactivated
vaccine against border disease pestivirus infection in sheep. In: Proceedings of the Second Symposium on
Pestiviruses, Edwards S., ed. Fondation Marcel Merieux, Annecy, France, 1–3 October 1992, 257–259
CARLSSON U. (1991). Border disease in sheep caused by transmission of virus from cattle persistently infected
with bovine virus diarrhoea virus. Vet. Rec., 128, 145–147.
CHAPPUIS G., BRUN A., KATO F., DAUVERGNE M., REYNAUD G. & DURET C. (1986). Etudes serologiques et
immunologiques realisees a la suite de I’isolement d’un pestivirus dans un foyer ovina chez des moutons de
L’Aveyron. In: Pestiviroses des Ovins et des Bovins, Espinasse J. & Savey M. eds. Ste Françoise de Buiatrie,
Paris, France, 55, 66.
DEKKER A., WENSVOORT G. & TERPSTRA C. (1995). Six antigenic groups within the genus pestivirus as identified by
cross-neutralisation assays. Vet. Microbiol., 47, 317–329.
ENTRICAN G., DAND A. & NETTLETON P.F. (1994). A double monoclonal-antibody ELISA for detecting pestivirus
antigen in the blood of viraemic cattle and sheep. Vet. Microbiol., 43, 65–74.
FENTON A., SINCLAIR J.A., ENTRICAN G., HERRING J.A. & NETTLETON P.F. (1991). A monoclonal antibody capture
ELISA to detect antibody to border disease virus in sheep sera. Vet. Microbiol., 28, 327–333.
FULTON R.W., D’OFFAY J.M., SALIKI J.T., BURGE L.J., HELMAN R.G., CONFER A.W., BOLIN S.R. & RIDPATH J.F. (1999).
Nested reverse transcriptase-polymerase chain reaction (RT-PCR) for typing ruminant pestiviruses: bovine viral
diarrhea viruses and border disease virus. Can. J. Vet. Res., 63, 276–281.
MCGOLDRICK A., BENSAUDE E., IBATA G., SHARP G. & PATON D.J. (1999). Closed one-tube reverse transcription
nested polymerase chain reaction for the detection of pestiviral RNA with fluorescent probes. J. Virol. Methods,
79, 85–95.
NETTLETON P.F., GILMOUR J.S., HERRING J.A. & SINCLAIR J.A. (1992). The production and survival of lambs
persistently infected with border disease virus. Comp. Immunol. Microbiol. infect. Dis., 15, 179–188.
NETTLETON P.F., GILRAY J.A., RUSSO P. & DLISSI E. (1998). Border disease of sheep and goats Vet. Res., 29, 327–
340.
OGUZOGLU T.C., FLOEGEL-NIESMANN G., FREY H.R. & MOENNIG V. (2001). Differential diagnosis of classical swine
fever and border disease: seroepidemiological investigation of a pestivirus infection on a mixed sheep and swine
farm. Dtsch Tierarztl. Wochenschr., 108, 210–213.
PATON D.J., SANDS J.J., LOWINGS J.P., SMITH J.E., IBATA G. & EDWARDS S. (1995). A proposed division of the
pestivirus genus into subgroups using monoclonal antibodies, supported by cross-neutralization assays and
genetic sequencing. Vet. Res., 26, 92–109.
RIDPATH J.F. & BOLIN S.R. (1997). Comparison of the complete genomic sequence of the border disease virus,
BD31, to other pestiviruses. Virus Res., 50, 237–243.
SANDVIK T., PATON D.J & LOWINGS P.J. (1997). Detection and identification of ruminant and porcine pestiviruses by
nested amplification of the 5’ untranslated cDNA region. J. Virol. Methods, 64, 43–56.
THABTI F., FRONZAROLI L., DLISSI E., GUIBERT J.M., HAMMAMI S., PEPIN M. & RUSSO P. (2002). Experimental model of
border disease virus infection in lambs: comparative pathogenicity of pestiviruses isolated in France and Tunisia.
Vet. Res., 33, 35–45.
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THEIL H.-J., COLLETT M.S., GOULD E.A., HEINZ F.X., HOUGHTON M., MEYERS G., PURCELL R.H. & RICE C.M. (2005).
Flaviviridae. In: Virus Taxonomy. Eighth report of the International Committee on Taxonomy of Viruses, Fauquet
C.M., Mayo M.A., Maniloff J., Desselberger U. & Ball L.A., eds. Elsevier Academic Press, 981–998.
THUR B., HILBE M., STRASSER M. & EHRENSPERGER F. (1997). Immunohistochemical diagnosis of pestivirus infection
associated with bovine and ovine abortion and perinatal death. Am. J. Vet. Res., 58, 1371–1375.
VALDAZO-GONZALEZ B., ALVAREZ-MARTINEZ M. & SANDVIK T. (2007). Genetic and antigenic typing of border Disease
virus isolates in sheep from the Iberian peninsula. Vet. J., 174, 316–324.
VAN RIJN P.A., VAN GENNIP H.G.P., LEENCLERSE C.H., BRUSCHKE C.J.M., PATON D.J., MOORMANN R.J.M. & VAN
OIRSCHOT J.T. (1997). Subdivision of the pestivirus genus based on envelope glycoprotein E2 Virology, 237, 337–
348.
VANTSIS J.T., BARLOW R.M., FRASER J. & MOULD D.L. (1976). Experiments in border disease VIII. Propagation and
properties of a cytopathic virus. J. Comp. Pathol., 86, 111–120.
VANTSIS J.T., RENNIE J.C., GARDINER A.C., WELLS P.W., BARLOW R.M. & MARTIN W.B. (1980). Immunisation against
Border disease. J. Comp. Path., 90, 349–354.
VILCEK S. (2001). Identification of pestiviruses contaminating cell lines and fetal calf sera. Acta Virol., 45, 81–86.
VILCEK S., HERRING A.J., HERRING J.A., NETTLETON P.F., LOWINGS J.P.L. & PATON D.J (1994) Pestiviruses isolated
from pigs, cattle and sheep can be allocated into at least three genogroups using polymerase chain reaction and
restriction endonucelase analysis. Arch. Virol., 136, 309–323.
VILCEK S. & NETTLETON P.F. (2006). Pestiviruses in wild animals Vet. Microbiol., 116, 1–12.
VILCEK S., NETTLETON P.F., PATON D.J. & BELAK S. (1997). Molecular characterization of ovine pestiviruses. J.
Gen. Virol., 78, 725–735.
VILCEK S. & PATON D.J. (2000). A RT-PCR assay for the rapid recognition of border disease virus. Vet. Res., 31,
437–445.
WENSVOORT G. & TERPSTRA C. (1988). Bovine viral diarrhoea virus infection in piglets born to sows vaccinated
against swine fever with contaminated virus. Res. Vet. Sci., 45, 143–148.
WENSVOORT G., TERPSTRA C. & DE KLUYVER E.P. (1989). Characterisation of porcine and some ruminant
pestiviruses by cross-neutralisation. Vet. Microbiol., 20, 291–306.
WILLOUGHBY K., VALDAZO-GONZALEZ, B., MALEY M., GILRAY J. & NETTLETON P.F. (2006). Development of a real time
RT-PCR to detect and type ovine pestiviruses. J. Virol. Methods, 132, 187–194.
*
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CHAPTER 2.7.2.
CAPRINE AND OVINE BRUCELLOSIS
(excluding Brucella ovis)
SUMMARY
Brucella melitensis (biovars 1, 2 or 3) is the main causative agent of caprine and ovine brucellosis.
Sporadic cases caused by B. abortus have been observed, but cases of natural infection are rare in
sheep and goats. Brucella melitensis is endemic in the Mediterranean region, but infection is
widespread world-wide. North America (except Mexico) is believed to be free from the agent, as are
Northern and Central Europe, South-East Asia, Australia and New Zealand.
Clinically, the disease is characterised by one or more of the following signs: abortion, retained
placenta, orchitis, epididymitis and, rarely, arthritis, with excretion of the organisms in uterine
discharges and in milk. Diagnosis depends on the isolation of Brucella from abortion material, udder
secretions or from tissues removed at post-mortem. Presumptive diagnosis of Brucella infection can
be made by assessing specific cell-mediated or serological responses to Brucella antigens.
Brucella melitensis is highly pathogenic for humans, causing Malta fever, one of the most serious
zoonoses in the world. All infected tissues, cultures and potentially contaminated materials should
therefore be handled at containment level 3.
Identification of the agent: Presumptive evidence of Brucella is provided by the demonstration, by
modified acid-fast staining of organisms typical of Brucella in abortion material or vaginal discharge,
especially if supported by serological tests. The polymerase chain reaction (PCR) methods provide
additional means of detection. Whenever possible, Brucella spp. should be isolated using selective
or non-selective media by culture from uterine discharges, aborted fetuses, udder secretions or
selected tissues, such as lymph nodes, spleen, uterus, testes and epididymes. Species and biovars
should be identified by phage lysis, and by cultural, biochemical and serological criteria. Molecular
methods have been developed that could also be used for complementary identification based on
specific genomic sequences.
Serological and allergic skin tests: The buffered Brucella antigen tests (BBAT) and the
complement fixation test (CFT) are usually recommended for screening flocks and individual
animals. The serum agglutination test is not considered to be reliable for use in small ruminants.
The indirect enzyme-linked immunosorbent assay (I-ELISA) and fluorescence polarisation assay
(FPA) can also be used for screening purposes. For pooled samples, there are no useful tests such
as the milk ring test for cattle. The brucellin allergic skin test can be used as a screening or
complementary test in unvaccinated flocks, provided that a purified, lipopolysaccharide (LPS)-free,
standardised antigen preparation is used. Results must then be interpreted in relation to the clinical
signs, history, and results of serological or cultural examination.
Requirements for vaccines and diagnostic biologicals: Brucella melitensis strain Rev.1 remains
the reference vaccine to immunise sheep and goats at risk of infection from B. melitensis and is the
vaccine with which any other vaccines should be compared. Production of Brucella antigens or
Rev.1 vaccine is based on a seed-lot system. Seed cultures to be used for antigens for serological
and allergic skin tests and for vaccines should originate from reference centres. They must conform
to minimal standards for viability, smoothness, residual infectivity and immunogenicity, purity,
identity and safety, if applicable. Brucellin preparations for the intradermal test must be free of
smooth lipopolysaccharide and must not produce nonspecific inflammatory reactions or interfere
with serological tests. Antigens for BBAT and CFT must be prepared from smooth strains of
B. abortus, strain 1119-3 or strain 99. Antigens for I-ELISA are prepared from B. abortus strain
1119-3 or strain 99 or B. melitensis biovar 1 reference strain 16M or antigens prepared from
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different smooth Brucella strains. All antigens must comply with minimum standards for purity,
sensitivity and specificity.
A. INTRODUCTION
Brucellosis in sheep and goats (excluding Brucella ovis infection) is primarily caused by one of the three biovars
of B. melitensis. Sporadic infections caused by B. abortus or B. suis have been observed in sheep and goats, but
such cases are rare. Pathologically and epidemiologically, B. melitensis infection in sheep and goats is very
similar to B. abortus infection in cattle (see Chapter 2.4.3 Bovine brucellosis). In most circumstances, the primary
route of transmission of Brucella is the placenta, fetal fluids and vaginal discharges expelled by infected ewes and
goats when they abort or have a full-term parturition. Shedding of Brucella is also common in udder secretions
and semen, and Brucella may be isolated from various tissues, such as lymph nodes from the head, spleen and
organs associated with reproduction (uterus, epididymides and testes), and from arthritic lesions (Alton et al.,
1988).
Brucella melitensis infection in domestic and wild susceptible species (see chapter 2.4.3) is not rare when these
species are reared in close contact with sheep and goats in enzootic areas. The manifestations of brucellosis in
these animals are similar to those in cattle or sheep and goats.
The World Health Organization (WHO) laboratory biosafety manual classifies Brucella (and particularly
B. melitensis) in Risk group III. Brucellosis is readily transmissible to humans, causing acute febrile illness –
undulant fever – which may progress to a more chronic form and can also produce serious complications affecting
the musculo–skeletal, cardiovascular, and central nervous systems. Infection is often due to occupational
exposure and is essentially acquired by the oral, respiratory, or conjunctival routes, but ingestion of dairy products
constitutes the main risk to the general public. There is an occupational risk to veterinarians, abattoir workers and
farmers who handle infected animals and aborted fetuses or placentas. Brucellosis is one of the most easily
acquired laboratory infections, and strict safety precautions should be observed when handling cultures and
heavily infected samples, such as products of abortion. Specific recommendations have been made for the safety
precautions to be observed with Brucella-infected materials (for further details see Chapter 1.1.3 Biosafety and
biosecurity in the veterinary microbiology laboratory and animal facilities, and refs 1, 39, 94 and 95 of chapter
2.4.3). Laboratory manipulation of live cultures or contaminated material from infected animals is hazardous, as is
handling large volumes of Brucella, and must be done under containment level 3 or higher conditions, as outlined
in chapter 1.1.3, to minimise occupational exposure.
The classification, microbiological and serological properties of the genus Brucella and related species and
biovars are given in the chapter 2.4.3.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
Refer to chapter 2.4.3 for the detailed agent identification procedure for Brucella.
2.
Serological tests
In situations where bacteriological examination is not practicable, diagnosis of Brucella infection must often be
based on serological methods (Alton et al., 1988; MacMillan, 1990). In routine tests, anti-Brucella antibodies are
detected in serum. The most widely used serum-testing procedures for the diagnosis of smooth Brucella
infections in sheep and goats are the buffered Brucella antigen tests (BBAT), and the complement fixation test
(CFT). The bulk milk ring test, which has been very useful in cattle, is ineffective in small ruminants.
In small ruminants, the BBAT and the CFT are the most widely used methods (Joint FAO/WHO Expert Committee
on Brucellosis, 1986). The indirect enzyme-linked immunosorbent assay (I-ELISA) and the fluorescence
polarisation assay (FPA) have shown similar diagnostic performance. All these tests are prescribed for
international trade. The BBAT is not completely specific, but is adequate as a screening test for detecting infected
flocks or for guaranteeing the absence of infection in brucellosis-free flocks. However, due to the relative lack of
sensitivity of both tests, discrepancies between results obtained using the Rose Bengal test (RBT) and the CFT
are not rare in infected sheep and goats (Blasco et al., 1994). The results of the two tests should therefore be
considered simultaneously to increase the likelihood of detecting infected individuals and to improve control of the
disease in areas where it has not been completely eradicated (Alton, 1990; Blasco, 1992; Blasco et al., 1994).
When, for practical or economic reasons, the CFT cannot be used simultaneously with the RBT in eradication
programmes, it is recommended to improve the sensitivity of the RBT by using three volumes of serum and one
volume of antigen (e.g. 75 µl and 25 µl, respectively) in place of an equal volume of each. This simple
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Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
modification increases RBT sensitivity and minimises the discrepancies between RBT and CFT results (Blasco et
al., 1994). Because antibodies induced after Rev.1 vaccination cannot be differentiated in both tests from those
induced by B. melitensis infection, RBT and CFT results should be carefully interpreted according to the
vaccination status in the flock. In addition, both tests are not specific enough to discriminate serological reactions
due to B. melitensis from the false-positive reactions (FPSR) due to cross-reacting bacteria such as Yersinia
enterocolitica O:9.
Good diagnostic results have been obtained in sheep and goats with indirect (I-) or competitive (C-) enzymelinked immunosorbent assays (ELISAs) using various antigens, but generally the ELISAs that use antigens with a
high content of smooth lipopolysaccharide (sLPS) are the most useful. The C-ELISA provides similar sensitivity to
the classical tests, RBT and CFT, and the I-ELISA has greater sensitivity. Like these classical tests, both ELISAs
are unable to differentiate B.-melitensis-infected animals from those recently vaccinated with the Rev.1 vaccine
(Marin et al., 1999) or infected with cross-reacting bacteria. Some of these ELISAs have potential advantages in
sensitivity and/or specificity with respect to both BBAT and CFT (Garin-Bastuji et al., 1998). Preliminary C-ELISAs
studies with a periplasmic protein from B. abortus (Rossetti et al., 1996) or B. melitensis (Cloeckaert et al., 1996)
as antigen have been applied in sheep and reported to be promising in differentiating Rev.1 vaccinated from
B. melitensis infected animals (Cloeckaert et al., 2001; Debbarh et al., 1996).

Reference sera
The OIE reference standards are those against which all other standards are compared and calibrated. For the
BBAT and CFT, please refer to chapter 2.4.3 for antigen standardisation and test protocols. A caprine reference
standard for ELISAs and FPA for sheep and goat antibodies has been developed and will be available to national
reference laboratories soon1.

Production of cells
Please refer to chapter 2.4.3. Brucella abortus biovar 1 strains 99 or 1119 are the only strains recommended for
the preparation of BBAT and CFT in sheep and goats.
a)
Brucella-buffered antigen test (a prescribed test for international trade)
Please refer to Chapter 2.4.3 Bovine brucellosis.

Antigen production
Please refer to chapter 2.4.3. Note that RB antigen made with B. abortus is usually used to test for
B. melitensis. The standardisation of RB antigen, as it is prescribed in chapter 2.4.3, provides a sufficient
sensitivity to the BBAT for international trade purposes. Moreover, it helps assure an adequate specificity in
free areas where FPSR occur because of cross-reacting bacteria such as Yersinia enterocolitica O:9.
However this standardisation is probably the main cause of the reduced sensitivity of some RB antigen
batches and of the discrepancies with the CFT (Blasco et al., 1994). Therefore, when RBT is used in
eradication programmes in endemic areas, it could be advisable to adjust the RB antigen titre so that it is
positive at a 1/45 OIEISS dilution and negative at a 1/55 dilution, without affecting the specificity of the test.
The discrepancies with the CFT can also be minimised by using three volumes of serum and one volume of
antigen (e.g. 75 µl and 25 µl, respectively) in place of an equal volume of each as mentioned in the standard
test procedure.

Test procedure
Please refer to chapter 2.4.3.
b)
Complement fixation test (a prescribed test for international trade)
•
Antigen production
Please refer to chapter 2.4.3. Note that CF antigen made with B. abortus is used to test for B. melitensis.

Test procedure
Please refer to chapter 2.4.3.
1
970
Obtainable from the OIE Reference Laboratory for Brucellosis at Animal Health and Veterinary Laboratories Agency
(AHVLA) Weybridge, New Haw, Addlestone, Surrey KT15 3NB, United Kingdom.
OIE Terrestrial Manual 2012
Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
c)
Enzyme-linked immunosorbent assays (a prescribed test for international trade)
Several variations of the I-ELISA have been described using different antigen preparations, antiglobulinenzyme conjugates, and substrate/chromogens. Several commercial I-ELISAs are available but before being
used for international trade, their respective cut-off should have been properly established using the
appropriate validation techniques (see Chapter 1.1.5 Principles and methods of validation of diagnostic assays
for infectious diseases) and these tests should be standardised against the above-mentioned Standard.
The test method is described in chapter 2.4.3.
d)
Fluorescence polarisation assay (a prescribed test for international trade)
The FPA for detection of caprine and ovine antibody to Brucella sp. is essentially the same as that described
for cattle (for more details see chapter 2.4.3); an example serum dilution used is 1/25 for the tube test and
1/10 for the plate test (Nielson et al., 1999; 2004; 2005; Ramirez-Pfeiffer et al., 2006). It is a simple
technique for measuring antigen/antibody interaction. The FPA may be used as a screening and/or
confirmatory test. Before being used for international trade, the FPA cut-off should be properly established
using the appropriate validation techniques (see chapter 1.1.5) and the test should be standardised against
the above-mentioned Standard.
3.
Other tests
a)
Brucellin skin test (an alternative test for international trade)
An alternative diagnostic test is the brucellin skin test, which can be used for screening unvaccinated flocks,
provided that a purified (free of sLPS) and standardised antigen preparation (e.g. brucellin INRA) is used.
The brucellin skin test has a high sensitivity for the diagnosis of B. melitensis infection in small ruminants
and, in absence of vaccination, is considered one of the most specific diagnostic tests (Alton et al., 1988;
Blasco, 1992; Garin-Bastuji et al., 1998; Joint FAO/WHO Expert Committee on Brucellosis, 1986). This test
is of particular value for the interpretation of FPSR due to infection with cross-reacting bacteria (FPSR
affected animals are always negative in the skin test), especially in brucellosis-free areas.
Rev.1 vaccinated animals can react in this test for years (Garin-Bastuji et al., 1998). Therefore this test
cannot be recommended either as the sole diagnostic test or for the purposes of international trade in areas
where Rev.1 vaccine is used.
To obtain suitable results it is essential to use standardised brucellin preparations that do not contain sLPS,
as this antigen may provoke antibody-mediated inflammatory reactions or induce antibodies that interfere
with subsequent serological screening. One such preparation is brucellin INRA, which is prepared from a
rough strain of B. melitensis that is commercially available2.

Test procedure
i)
A volume of 0.1 ml of brucellin is injected intradermally into the lower eyelid.
ii)
The test is read after 48 hours.
iii)
Any visible or palpable reaction of hypersensitivity, such as an oedematous reaction leading to an
elevation of the skin or thickening of the eyelid (≥ 2 mm), should be interpreted as a positive reaction.
Although in the absence of vaccination the brucellin intradermal test is one of the most specific tests in
brucellosis, diagnosis should not be made exclusively on the basis of positive intradermal reactions and
should be supported by adequate serological tests. The intradermal inoculation of brucellin might induce a
temporary anergy in the cellular immune response. Therefore an interval of 6 weeks is generally
recommended between two tests repeated on the same animal.
b)
Native hapten tests
The native hapten-based gel precipitation tests3 (as described in chapter 2.4.3.) are also of interest in sheep
and goats as they are very specific for discriminating the serological responses of infected animals (positive)
from those induced in Rev.1 vaccinated animals (usually negative after a given time after vaccination).
2
3
Brucellergène OCB®, Synbiotics Europe, 2 rue Alexander Fleming, 69007 Lyon, France.
The detailed procedure could be obtained from the Departamento de Sanidad Animal, Centro de Investigacion y
Tecnologia Agroalimentaria/Gobierno de Aragon, Avenida Montañana 930, 50059. Zaragoza. Spain.
OIE Terrestrial Manual 2012
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Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
The optimal diagnostic sensitivity (around 90%) is obtained in the double gel diffusion (DGD) or reverse
radial immunodiffusion tests for sheep and goats, respectively (Debbarh et al., 1996; Joint FAO/WHO Expert
Committee on Brucellosis, 1986).
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
C1. Brucellin
Please refer to chapter 2.4.3.
C2. Vaccines
Brucella melitensis strain Rev.1 vaccine
The most widely used vaccine for the prevention of brucellosis in sheep and goats is the Brucella melitensis
Rev.1 vaccine, which remains the reference vaccine with which any other vaccines should be compared. The
RB51 vaccine is not effective in sheep against B. melitensis infection (El-Idrissi et al., 2001). In addition, other
rough mutants defective in core and O-polysaccharide synthesis and export induce antibodies reacting in the IELISA with sLPS and are less effective than Rev.1 vaccine against B. melitensis infection in sheep (Barrio et al.,
2009). The Rev.1 vaccine is used as a freeze-dried suspension of live B. melitensis biovar 1 Rev.1 strain for the
immunisation of sheep and goats. It should be normally given to lambs and kids aged between 3 and 6 months as
a single subcutaneous or conjunctival inoculation. The standard dose is between 0.5 × 109 and 2.0 × 109 viable
organisms. The subcutaneous vaccination induces strong interferences in serological tests and should not be
recommended in combined eradication programmes (Díaz-Aparicio et al., 1994; Marin et al., 1999). However,
when this vaccine is administered conjunctivally, it produces a similar protection without inducing a persistent
antibody response, thus facilitating the application of eradication programmes combined with vaccination (DíazAparicio et al., 1994; Marin et al., 1999). Care must be taken when using Rev.1 vaccine to avoid the risk of
contaminating the environment or causing human infection. In many developing countries and endemic areas,
vaccination of the whole population has to be considered as the best option for the control of the disease (Blasco,
1997). However, Rev.1 vaccine is known to often cause abortion and excretion in milk when animals are
vaccinated during pregnancy, either with a full or reduced dose (Blasco, 1997). These side-effects are
considerably reduced when adult animals are vaccinated conjunctivally (full dose) before mating or during the last
month of pregnancy. Therefore, when mass vaccination is the only means of controlling the disease, a
vaccination campaign should be recommended using the standard dose of Rev.1 administered by the conjunctival
route when the animals are not pregnant or during the late lambing and prebreeding season (Blasco, 1997).
The subcutaneous vaccination of young animals and the vaccination of adult animals, even at reduced doses,
may lead to long-term persistence of vaccinal antibodies in a significant proportion of vaccinated animals that
creates serious interferences in the serological diagnosis of brucellosis. As indicated above, conjunctival
vaccination minimises these problems and thus it is the recommended method for combined eradication
programmes. Therefore, the serological diagnosis of brucellosis should take into account the vaccinal state of the
herd and the overall frequency distribution of antibody titres detected in the group of animals tested.
1.
Seed management
a)
Characteristics of the seed
Brucella melitensis biovar 1 strain Rev.1 original seed for vaccine production can be obtained commercially4.
A European reference Rev.1 strain that possesses the characteristics of the Rev.1 original seed is also
obtainable from the European Pharmacopoeia5.
Production of Brucella live vaccines is based on the seed-lot system described above (Section B.2) for
BBAT and CFT antigens. Strains should be cultured in a suitable medium. Strain Rev.1 must conform to the
characteristics of B. melitensis biovar 1, except that it should grow more slowly. Additionally, when incubated
in air (atmospheres containing CO2 alter the results) at 37°C, it should grow on agar containing streptomycin
(2.5 µg/ml), and it should be inhibited by the addition to a suitable culture medium of sodium benzylpenicillin
(3 µg [5 International Units (IU)]/ml), thionin (20 µg/ml) or basic fuchsin (20 µg/ml). Recently, polymerase
chain reaction and molecular techniques have been used to further characterise the vaccine (Bardenstein et
al., 2002; Cloeckaert et al., 2002). It must also conform to the characteristics of residual virulence and
immunogenicity in mice of the original seed.
4
5
972
Obtainable from the OIE Reference Laboratory for Brucellosis at Anses Maisons-Alfort, 94706, France.
Obtainable from the European Pharmacopoeia, BP 907, 67029 Strasbourg Cedex 1, France.
OIE Terrestrial Manual 2012
Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
b)
Method of culture
Serum–dextrose agar, and trypticase–soy agar, to which 5% serum or 0.1% yeast extract may be added,
are among the solid media that have been found to be satisfactory for propagating the Rev.1 strain (Alton et
al., 1988; WHO, 1977). Rev.1 strain does not grow well on potato agar.
For vaccine production, Rev.1 may be grown under conditions similar to those described for S99 and S1119-3
(see chapter 2.4.3), except that Rev.1 generally needs 3–5 days to grow, the phenol saline is replaced by a
freeze-drying stabiliser, and the organisms are not killed but are stored at 4°C while quality control examinations
are carried out as described below. Moreover, the specific requirements for Rev.1 vaccine production recommend
that: each seed lot (i.e. the culture used to inoculate medium for vaccine production) should be no more than three
passages removed from an original seed culture and that the harvest of a vaccine lot should be no more than
three passages from a seed lot or an original seed. The original seed culture should always be checked for the
absence of dissociation before use. The recommended method for preparing seed material is given in ref. 2. The
following freeze-drying stabiliser (sterilised by filtration) is of proven value: enzymatic digest of casein (2.5 g);
sucrose (5 g); sodium glutamate (1 g); distilled water (100 ml).
c)
Validation as a vaccine
Numerous independent studies have confirmed the value of B. melitensis strain Rev.1 as a vaccine for
protecting sheep and goats from brucellosis. Its virulence is unchanged after passage through pregnant
sheep and goats. Abortions may result when the Rev.1 vaccine is inoculated into pregnant ewes or goats.
The vaccine-induced abortions are not avoided using reduced doses, and doses as low as 106, used either
subcutaneously or conjunctivally, have been demonstrated to induce abortions and milk excretion of the
vaccine strain (Blasco, 1997).
A Rev.1 vaccine is efficient if it possesses the characteristics of the Rev.1 original strain, i.e. those of
B. melitensis biovar 1 reference strain 16M (ATCC No. 23456), except those specific for the strain Rev.1
(Alton et al., 1988; Joint FAO/WHO Expert Committee on Brucellosis, 1986), and if it proves to be
satisfactory with respect to immunogenicity and residual virulence in the mouse model (Bosseray, 1992)
(see below).
2.
Method of manufacture (Alton et al., 1988; WHO, 1977)
For production of B. melitensis strain Rev.1 vaccine, the procedures described above for antigens (Alton et al.,
1988) can be used except that the cells are collected in a freeze-drying stabiliser and deposited by centrifugation.
The yield from one fermenter run or the pooled cells from a batch of Roux flask cultures inoculated on the same
occasion from the same seed lot constitutes a single harvest. More than one single harvest may be pooled to
form the final bulk that is used to fill the final containers of a batch of vaccine. Before pooling, each single harvest
must be checked for purity, cell concentration, dissociation and identity. The volume of the final bulk is adjusted
by adding sufficient stabiliser so that a dose contains an appropriate number of viable organisms. After adjusting
the cell concentration of the final bulk, tests for identity, dissociation and absence of contaminating organisms are
conducted (see below).
3.
In-process control
In-process checks should be made on the growth of Rev.1 vaccine from either solid or liquid medium to verify
identity and to ensure purity and freedom from dissociation to rough forms during preparation of seed lots, single
harvests, final bulks and the final (filling) lots. At least 99% of cells in seed lots and 95% of cells in final lots should
be in the smooth phase.
Cell concentration should be estimated on the bulks and precisely determined on final lots. Immunogenicity and
the residual virulence (50% persistence time or 50% recovery time) should also be determined on seed lots and
final lots. If these tests have been done with good results on a representative batch of the test vaccine, it does not
have to be repeated routinely on other vaccine lots prepared from the same seed lot and with the same
manufacturing process.
4.
Batch control
With freeze-dried vaccine, the control tests should be conducted on the vaccine reconstituted in the form in which
it will be used.
a)
Sterility (or absence of extraneous microorganisms)
The Rev.1 vaccine should be checked for bacterial and fungal contamination as prescribed in Chapter 1.1.7
Tests for sterility and freedom from contamination of biological materials.
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Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
b)
Safety
The Rev.1 vaccine is a virulent product per se, and it should keep a minimal virulence to be efficacious (see
Section C2.4.c in chapter 2.4.3).
c)
Potency
A Rev.1 vaccine is efficient if it possesses the characteristics of the Rev.1 original strain, i.e. if it is
satisfactory with respect to immunogenicity, and residual virulence (Bosseray, 1993). Batches should also
be checked for the number of viable organisms.

Identity
The reconstituted Rev.1 vaccine should not contain extraneous microorganisms. Brucella melitensis
present in the vaccine is identified by suitable morphological, serological and biochemical tests and by
culture: when incubated in air at 37°C, Rev.1 strain is inhibited by addition to the suitable culture
medium of 3 µg (5 IU) per ml of sodium benzyl-penicillin, thionin (20 µg/ml) or basic fuchsin (20 µg/ml);
the strain grows on agar containing 2.5 µg per ml of streptomycin.

Smoothness (determination of dissociation phase)
Please refer to chapter 2.4.3.
Sometimes slight and difficult to observe differences, can be seen in the size of Rev.1 colonies. The
small colonies (1–1.2 mm in diameter) are typical for Rev.1, but larger Rev.1 colonies can appear
depending on the medium used, the amount of residual moisture in the incubator atmosphere, and the
presence or absence of CO2. The frequency of variation in colony size occurs normally at a ratio of
1 large to 103 small colonies. Both Rev.1 variants are of the S (smooth) type. To avoid an increase in
this colony size variation along successive passages, it is important to always select small colonies for
preparation of seed lots.

Enumeration of live bacteria
Please refer to chapter 2.4.3.

Residual virulence (50% persistence time or 50% recovery time) (Bosseray, 1991; Grillo et al., 2000)
The same technical procedures indicated for 50% recovery time (RT50) calculation of S19 vaccine (see
chapter 2.4.3) have to be applied for Rev.1, except that B. abortus S19 seed lot or batch to be tested
(test vaccine) and the S19 original seed culture (used as a reference strain), respectively, are replaced
by the corresponding B. melitensis Rev.1 seed lot or batch to be tested (test vaccine) and the
B. melitensis Rev.1 original seed culture as the reference strain. For the reference original Rev.1 strain,
RT50 and confidence limits are around 7.9 ± 1.2 weeks. A given Rev.1 vaccine seed lot or batch should
keep similar residual virulence to be acceptable.
If this test has been done with good results on a representative batch of the test vaccine, it does not have to
be repeated routinely on other vaccine lots prepared from the same seed lot and with the same
manufacturing process.

Immunogenicity in mice
The same technical procedures indicated for immunogenicity calculation of S19 vaccine (see chapter
2.4.3) have to be applied for Rev.1, except that B. abortus S19 seed lot or batch to be tested (test
vaccine) and the B. abortus S19 original seed culture (used as a reference strain), respectively, are
replaced by the corresponding B. melitensis Rev.1 seed lot or batch to be tested (test vaccine) and the
B. melitensis Rev.1 original seed culture as the reference strain.
Conditions of the control experiment are satisfactory when: i) the response in unvaccinated mice (mean
of Y) is at least of 4.5; ii) the response in mice vaccinated with the reference Rev.1 vaccine is lower
than 2.5; and iii) the standard deviation calculated on each lot of six mice is lower than 0.8.
If this test has been done with good results on a representative batch of the test vaccine, it does not have to
be repeated routinely on other vaccine lots prepared from the same seed lot and with the same
manufacturing process.
d)
Duration of immunity
It is accepted that subcutaneous or conjunctival vaccination with standard doses of Rev.1 confers a solid
and durable immunity in sheep and goats. However, growing field evidence shows that the immunity
conferred declines with time, and revaccination could be advisable in endemic areas.
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Chapter 2.7.2. — Caprine and ovine brucellosis (excluding Brucella ovis)
The use of reduced doses of Rev.1 produces a less efficient immunity, while side-effects, such as antibody
responses or induction of abortion, are not fully avoided.
e)
Stability
Strain Rev.1 vaccine prepared from seed stock from appropriate sources is stable in characteristics provided
that the in-process and batch control requirements described above are fulfilled, and shows no tendency to
reversion to virulence. The lyophilised vaccine shows a gradual loss of viable count, but should retain its
potency for the recommended shelf life. Allowance for this phenomenon is normally made by ensuring that
the viable count immediately following lyophilisation is well in excess of the minimum requirement.
Maintenance of a cold chain during distribution of the vaccine will ensure its viability.
f)
Preservatives
Antimicrobial preservatives must not be used in live Rev.1 vaccine. For preparation of the freeze-dried
vaccine, a stabiliser as described in Section C2.4.f of chapter 2.4.3 is recommended.
g)
Precautions (hazards)
Please refer to chapter 2.4.3. Brucella melitensis Rev.1, although an attenuated strain, is still capable of
causing disease in humans. Accordingly, cell cultures and suspensions must be handled under appropriate
conditions of biohazard containment (see chapter 1.1.3). Reconstitution and subsequent handling of the
reconstituted vaccine should be done with care to avoid accidental injection or eye or skin contamination.
Vaccine residues and injection equipment should be decontaminated with a suitable disinfectant. Medical
advice should be sought in the event of accidental exposure. The efficacy of the antibiotic treatment of
infections caused by Rev.1 (a streptomycin-resistant strain) in humans has not been adequately established
but data in mice suggest that if Rev.1 contamination occurs, a combined treatment with doxycycline plus
rifampicin or gentamycin could be recommended (Grillo et al., 2006).
5.
Tests on the final product
a)
Safety
See Section C2.4.b of chapter 2.4.3.
b)
Potency
For the freeze-dried vaccine, the potency must be determined on the final product. The tests are as
described in Section C2.4.c of chapter 2.4.3.
In order to assess the vaccine efficiency, a representative sample of previously seronegative animals
vaccinated with each new vaccine batch should be bled 15–20 days after vaccination and the serum
samples submitted to BBAT. If adequate and independently of the vaccination route used, more than 80% of
vaccinated animals should be BBAT positive.
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LETESSON J.J., GORVEL J.P., MORIYÓN I., BLASCO J.M. & ZYGMUNT M.S. (2009). Rough mutants defective in core and
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immunogenicity in mice. Biologicals, 19, 355–363.
BOSSERAY N. (1992). Le vaccin Rev.1: dérive des caractères d’immunogénicité et de virulence indépendante des
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BOSSERAY N. (1993). Control methods and thresholds of acceptability for anti-Brucella vaccines. Dev. Biol. Stand.,
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CLOECKAERT A., BAUCHERON S., VIZCAINO N. & ZYGMUNT M.S. (2001). Use of recombinant BP26 protein in
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of fluorescence polarization assay with card and complement fixation tests for the diagnosis of goat brucellosis in
a high prevalence area. Vet. Immunol. Immunopathol., 110, 121–127.
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WORLD HEALTH ORGANIZATION (1977). Requirements for Brucella melitensis strain Rev.1 vaccine (Live – for
veterinary use). World Health Organization (WHO) Technical Report Series No. 610, 28th Report, Annex 4. WHO,
Geneva, Switzerland, 85–97.
*
* *
NB: There are OIE Reference Laboratories for Caprine and ovine brucellosis (excluding Brucella ovis)
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for caprine and ovine brucellosis (excluding Brucella ovis)
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CHAPTER 2.7.3/4.
CAPRINE ARTHRITIS-ENCEPHALITIS
& MAEDI-VISNA
SUMMARY
Maedi-visna (MV) and caprine arthritis-encephalitis (CAE) are persistent lentivirus infections of
sheep and goats and are often grouped together as the small ruminant lentiviruses (SRLVs).
Maedi-visna is also known to as ovine progressive pneumonia (OPP). Phylogenetic analyses
comparing nucleotide sequences of MV virus (MVV) and CAE virus (CAEV) has demonstrated that
these are closely related lentiviruses. One source of CAEV and MVV transmission is colostrum and
milk. The source of horizontal transmission in the absence of lactation remains unknown; however,
faeces and lung fluids are known to harbour infectious virus. Ovine lentiviruses have been identified
in most of the sheep-rearing countries of the world, with the notable exceptions of Australia and
New Zealand. The distribution of CAEV is highest in industrialised countries, and seems to have
coincided with the international movement of European breeds of dairy goats. Clinical and
subclinical MV and CAE are associated with progressive, mononuclear cell inflammatory lesions in
the lungs, joints, udder and central nervous system. Indurative mastitis is common in both species,
and its economic significance may be underestimated. Laboured breathing associated with
emaciation caused by progressive pneumonitis is the predominant features in clinically affected
sheep, whereas polyarthritis is the main clinical sign in goats. However, most lentivirus-infected
sheep and goats are largely asymptomatic, but remain persistent carriers of virus and are capable
of transmitting infection via colostrum or milk and respiratory secretions. The most practical and
reliable approach to confirming a diagnosis of MV or CAE is a combination of serology and clinical
evaluation. Although serology represents the most cost-effective method of diagnosing persistently
infected, clinically normal animals, it should be understood that testing errors occur. The frequency
of error depends on several factors including but not limited to: 1) the assay format, 2) the
homology between the strain of virus used in the assay and the strains of virus present in the
testing populations, and 3) the viral antigen used in the assay.
Identification of the agent: Virus isolation can be attempted from live clinical or subclinical cases
by co-cultivating peripheral blood or milk leukocytes with appropriate ovine or caprine cell cultures,
such as choroïd plexus (MVV) or synovial membrane (CAEV) cells. Virus isolation is very specific
but has variable sensitivities. Following necropsy, virus isolation is most readily accomplished by
establishing explant cultures of affected tissues, e.g. lung, choroïd plexus, synovial membrane or
udder. Also, alveolar macrophages may be obtained from the lung at post-mortem and co-cultivated
with susceptible cells. The cytopathic effects are characteristic, consisting of the appearance of
refractile stellate cells and syncytia. The presence of MVV or CAEV can be confirmed by
immunolabelling methods and electron microscopy.
Nucleic acid recognition methods: Many standard and a few quantitative polymerase chain
reaction (PCR) assays for detecting MV and CAE provirus have been described and are now being
used routinely in many laboratories for the rapid detection, quantitation, and identification of the
small ruminant lentivirus strains. Cloning and/or sequencing of PCR products is the most direct
method to confirm specificity of PCR products.
Serological tests: Most infected sheep and goats possess detectable specific antibodies that can
be assayed by a number of different serological tests. The two most commonly used are the agar
gel immunodiffusion test and the enzyme-linked immunosorbent assay (ELISA). Western blot
analysis and radio-immunoprecipitation are also performed, but only in specialised laboratories. A
milk antibody assay may be appropriate in dairy goat herds. The time required for seroconversion
following infection can be relatively prolonged and unpredictable, being measured in months rather
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than in weeks. However, after seroconversion, the antibody response usually persists and antibodypositive sheep and goats are regarded as virus carriers.
Requirements for vaccines and diagnostic biologicals: There are no biological products
available.
A. INTRODUCTION
Maedi-visna (MV), of sheep and caprine arthritis/encephalitis (CAE) of goats are persistent virus infections
caused by closely related lentiviruses (Peterhans et al., 2004). Maedi-visna is also known as ovine progressive
pneumonia (OPP). Sheep can be experimentally infected with CAE and goats can be experimentally infected with
MV (Banks et al., 1983). In addition, phylogenetic analyses comparing nucleotide sequences of MV virus (MVV)
and CAE virus (CAEV) show clear indications of the existence and epidemiological importance of cross-species
transmission between sheep and goats without demonstrating clearly that one virus has emerged from the other
(Herrmann et al., 2004; Karr et al., 1996; Leroux et al., 1997; Roland et al., 2002; Shah et al., 2004a; 2004b;
Valas et al., 1997; Zanoni, 1998). MV and CAE are characterised by lifelong persistence of the causal agent in
host monocytes and macrophages, and a variable length of time between infection and induction of a
serologically detectable antiviral antibody response. Most infected sheep and goats do not exhibit clinical disease,
but remain persistently infected and are capable of transmitting virus (Adams et al., 1983; Cork, 1990; Crawford et
al., 1980).
Maedi-visna is an Icelandic name that describes two of the clinical syndromes recognised in MV virus (MVV)infected sheep. ‘Maedi’ means ‘laboured breathing’ and describes the disease associated with a progressive
interstitial pneumonitis, and ‘visna’ means ‘shrinkage’ or ‘wasting’, the signs associated with a paralysing
meningoencephalitis. Whereas progressive lung disease is the primary finding with MVV infection, chronic
polyarthritis, with synovitis and bursitis is the primary clinical outcome of CAE virus (CAEV) infection. Encephalitis
occurs primarily in kids aged between 2 and 6 months following CAEV infection, but careful differential diagnoses
need to be conducted to rule out other syndromes or infections in kids. Indurative mastitis occurs in both
syndromes. The lungs of sheep affected by MV do not collapse when removed from the thorax and often retain
the impression of the ribs. The lungs and lymph nodes increase in weight (up to 2–3 times the normal weight).
The lesions are uniformly distributed throughout the lungs, which are uniformly discoloured or mottled grey-brown
in colour and of a firm texture. Udders affected by MV are diffusely indurated and associated lymph nodes may be
enlarged.
When MV or CAE is the suspected cause of clinical disease, confirmation of the diagnosis can be achieved by a
combination of clinical evaluation, serology and, when necessary, histological examination of appropriate tissues
collected at necropsy. Important tissues to examine include lung for progressive interstitial pneumonitis, brain and
spinal cord for meningoencephalitis, udder for indurative mastitis, affected joints and synovium for arthritis, and
kidney for vasculitis (Crawford & Adams, 1981; Cutlip et al., 1985; 1986; Knowles et al., 1990; Oliver et al., 1981a;
1981b). The nature of the inflammatory reaction in each site is similar, consisting of an interstitial, mononuclear
cell reaction, sometimes with large aggregates of lymphoid cells and follicle formation.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
Isolation and characterisation of MVV or CAEV would not normally be attempted for routine diagnostic purposes.
Due to the persistent nature of these infections, the establishment of a positive antibody status is sufficient for the
identification of virus carriers. However, due to a late seroconversion after infection, negative serology may occur
in recently infected animals.
There are two approaches to the isolation of MVV and CAEV: one for use with the live animal, and the second for
use with necropsy tissues.
a)
Isolation from the live animal
•
Maedi-visna virus
The MV provirus DNA is carried in circulating monocytes and tissue macrophages. Virus isolation from the
live animal therefore requires the establishment of leukocyte preparations, with aseptic precautions, from
peripheral blood or milk during lactation, culturing them together with indicator cells. Sheep choroïd plexus
(SCP) cells are commonly used for this purpose. These indicator cells can be prepared as primary explant
cultures from fetal or newborn virus-free lambs, and their number can be multiplied over three to four
passages for storage in liquid nitrogen. The recovered SCP cells are suitable for co-cultivation for up to 10 or
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15 passages. Although the cells continue to grow well thereafter, their susceptibility to MVV may become
reduced.
Leukocyte preparations can be made from peripheral blood as buffy coats by the centrifugation at 1000 g of
heparinised, ethylene diamine tetra-acetic acid (EDTA) or citrated samples for 15 minutes. The cells are
aspirated, suspended in Hanks’ balanced salt solution (HBSS), and further purified by centrifugation at 400 g
on to a suitable cushion (e.g. Ficoll Paque [Pharmacia]) for 40 minutes. The interface cells are spin-washed
once or twice in HBSS at 100 g for 10 minutes, and the final cell pellet is resuspended in medium to a
concentration of approximately 106 cells/ml; cells are generally cultured for 10–12 days in Teflon bags and
are then added to a washed monolayer of slightly subconfluent SCP cells in a flask with an area of 25 cm2.
Leukocytes can be similarly deposited from milk by centrifugation, when they are spin-washed, resuspended
and finally added to SCP monolayer cultures.
These cultures are maintained at 37°C in a 5% CO2 atmosphere, changing the medium and passaging as
necessary. They are examined for evidence of a cytopathic effect (CPE), which is characterised by the
appearance of refractile stellate cells with dendritic processes accompanied by the formation of syncytia.
The cultures should be maintained for several weeks before being discarded as uninfected. Once a CPE is
suspected, cover-slip cultures should be prepared. These are fixed, and evidence of viral antigen is sought
by immunolabelling, usually by means of indirect fluorescent antibody or by the use of indirect
immunoperoxidase methods. In addition, the cells of any suspect monolayers are deposited by
centrifugation, and preparations are made for the identification of any characteristic lentivirus particles by
transmission electron microscopy. Reverse transcriptase in the supernatant of the cell culture is indicative of
the presence of retroviruses.
•
Caprine arthritis/encephalitis virus
The same principles that apply to the isolation of MVV also apply to the isolation of CAEV. CAEV was
originally isolated by explantation of synovial membrane from an arthritic goat (Crawford & Adams, 1981).
With live CAEV-infected goats, peripheral blood, milk, and possibly joint fluid aspirate represent the most
suitable specimens from which leukocyte preparations can be established. Goat synovial membrane (GSM)
cells are suitable indicator cells. If a CPE is suspected, tests for detection of viral antigen should be carried
out, as described above.
b)
Isolation from necropsy tissues
•
Caprine arthritis/encephalitis virus and Maedi-visna virus
Samples of suspect tissues, collected as fresh as possible, such as lung, synovial membranes, udder, etc.,
are collected aseptically into sterile HBSS or cell culture medium and minced finely in a Petri dish using
scalpel blades. Individual fragments are collected by Pasteur pipette and transferred to flasks of 25 cm2,
approximately 20–30 fragments per flask, and a drop of growth medium is placed carefully on each. The
flasks are then incubated at 37°C in a humid 5% CO2 atmosphere, and left undisturbed for a few days to
allow the individual explants to adhere to the plastic. Fresh medium can be added with care, after which rafts
of cells will gradually grow out from the fragments. When there is sufficient cell out-growth, the cultures are
trypsin dispersed to allow the development of cell monolayers. These can be examined for CPE, and any
suspected virus growth is confirmed in the same way as for the co-cultivations.
Adherent macrophage cultures are easy to establish from lung-rinse material (post-mortem broncho-alveolar
lavage) and can be tested for virus production by serology, electron microscopy, or reverse transcriptase
assay within 1–2 weeks. Virus isolations can be done by co-cultivation of macrophages and SCP or GSM
cells as described for leukocytes above.
c)
Nucleic acid recognition methods
Most virus disease diagnostic laboratories will be equipped for the basic cell culture procedures described
above. Many laboratories can now also perform nucleic acid recognition methods for the detection,
quantitation, and identification of MV and CAE proviral DNA using the standard polymerase chain reaction
(PCR) followed by Southern blotting, in situ hybridisation, or cloning and/or sequencing of the PCR product
(Alvarez et al., 2006; Herrmann-Hoesing et al., 2007; Johnson et al., 1992). Standard PCR techniques for
the detection of MV and CAE proviral DNA in cells and tissues are routinely used in many laboratories and
are generally used as supplemental tests for determining infection status of those animals that cannot be
definitively diagnosed by serology (Deandres et al., 2005; Knowles, 1997). Real-time or quantitative PCR
techniques are beginning to be used in a few laboratories and these tests, in addition to determining
infection status, also quantify the amount of MV or CAE provirus in an animal (Alvarez et al., 2006;
Herrmann-Hoesing et al., 2007). Furthermore, molecular techniques such as PCR, cloning and sequencing
also provide knowledge on a country’s or region’s specific MV and CAE strains, which may influence which
serological assay and corresponding MV or CAE antigen to use. Phylogenetic analyses of MV and CAE
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proviral DNAs from SRLV strains throughout the world have suggested that in some areas, MV may have
naturally infected goats, and CAE may have naturally infected sheep (Shah et al., 2004a; 2004b). In the
future, molecular diagnostic tests along with phylogenetic analyses of MV and CAE provirus may be used to
track transmission.
An important issue in the use of PCR is specificity. Because of the possibility of amplifying unrelated
sequences from the host’s genomic DNA (false positives), the amplified product should be checked by either
hybridisation, restriction endonuclease digestion patterns, or sequencing. Sequencing provides the best
proof of specificity in the validation of PCR-based tests and is recommended by the OIE. Sensitivity of PCRbased tests can be improved by the use of nested PCR, but specificity of the nested PCR test should be
checked using hybridisation, restriction endonuclease digestion patterns, or sequencing methods.
2.
Serological tests
Ovine and caprine lentivirus infections are persistent, so antibody detection is a valuable serological tool for
identifying virus carriers. The close antigenic relationship between MVV and CAEV does not extend to
detection of heterologous antibody in some serological assays (Knowles et al., 1994).
The assays most commonly used to serologically diagnose the presence of a small ruminant lentivirus
infection are agar gel immunodiffusion (AGID) and the enzyme-linked immunosorbent assay (ELISA). AGID
was first developed and reported in 1973 (Terpstra & De Boer, 1973), and the ELISA was first developed
and reported in 1982 (Houwers et al., 1982). The AGID is specific, reproducible and simple to perform, but
experience is required for reading the results. The ELISA is economical, quantitative and can be automated,
thus making it useful for screening large numbers of sera. The sensitivity and specificity of both the AGID
assay and ELISA depend upon the virus strain used in the assay, viral antigen preparation, and the
standard of comparison assay. Western blot analysis and/or radio-immunoprecipitation are the standards of
comparison used to access sensitivity and specificity of new AGID tests and ELISAs.
a)
Agar gel immunodiffusion (a prescribed test for international trade)
There are two MV and CAE viral antigens of major importance in routine serology, a viral surface envelope
glycoprotein commonly referred to as SU or gp135, and a nucleocapsid protein referred to as CA or p28.
These are both conserved in an antigen preparation consisting of medium harvested from infected cell
cultures and concentrated approximately 50-fold by dialysis against polyethylene glycol. As an example the
WLC-1 strain of MV virus is commonly used in the AGID assay in the United States (Cutlip et al., 1977)1 and
a Canadian MV field strain is used for AGID tests in Canada (Simard & Briscoe, 1990b).
It is important to recognise that the sensitivity of the AGID test for detecting anti-CAEV antibody is
dependent on both the virus strain and the viral antigen used (Adams & Gorham, 1986; Knowles et al.,
1994). It was demonstrated that an AGID test with CAEV gp135 afforded greater sensitivity than an AGID
test with CAEV p28 (Adams & Gorham, 1986). Also, it was shown that when compared with radioimmunoprecipitation, the sensitivity of the AGID test for anti-CAEV antibody was 35% greater using CAE
virus antigen over using MV virus antigen (Knowles et al., 1994). The most likely explanation for this
difference in sensitivity between the CAE and MV virus antigen for the detection of anti-CAEV antibody is
that although the radio-immunoprecipitation assay requires only the binding of a single epitope by antibody
to obtain a positive result, precipitation in an agar gel requires multiple epitope–antibody interactions.
Although the MV and CAE viruses have 73–74.4% nucleotide sequence identity in the envelope gene
(Herrmann et al., 2004), this amount of identity may not be sufficient to produce sufficient antibody to CAE
and MV mutually common epitopes resulting in undetectable antibody/antigen precipitin lines using MV virus
antigen. When the appropriate antigen is used, the AGID test performance is high. When compared with
immunoprecipitation, the AGID for the detection of anti-CAEV antibody, if CAEV antigen was used, had 92%
sensitivity and 100% specificity (Knowles et al., 1994). In addition, the AGID for detection of anti-MVV
antibody, if MVV antigen was used, had 99.3 and 99.4% sensitivity and specificity, respectively (Herrmann et
al., 2003b).
In adult persistently MVV-infected sheep and CAEV-infected goats, the predominant immunoprecipitating
antibody response is directed against gp135 antigen (Herrmann et al., 2005; Knowles et al., 1990). An antip28 response is usually present at a lower titre than the anti-gp135 response in persistently infected adult
small ruminants using immunoprecipitation. In some CAEV-infected goats there is evidence to suggest that
an anti-gp135 antibody response is produced, in the absence of an anti-p28 response and vice versa, in a
proportion of individuals (Dawson, 1985; Rimstad et al., 1994). Hence, for validation of a test, standard sera
producing both anti-gp135 and anti-p28 precipitin lines are required.
1
This virus has been distributed by Dr Howard Lehmkuhl, National Animal Disease Center, United States Department of
Agriculture, P.O. Box 70, Ames, Iowa, USA.
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The gel medium is 0.7–1% agarose in 0.05 M Tris buffer, pH 7.2, with 8.0% NaCl. The test is conveniently
performed in plastic Petri dishes, or in 10 cm2 plastic trays. The pattern and size of the wells will determine
the number of sera tested per plate. Various well patterns can be adopted, but a hexagonal arrangement
with a central well is usual: for example, a pattern with alternating large (5 mm in diameter) and small (3 mm
in diameter) peripheral wells, 2 mm apart and 2 mm from a central antigen well that is 3 mm in diameter. The
large peripheral wells are used for test sera and the small ones for standard sera. A weak positive control
must be included in each test. The plates are incubated overnight at 20–25°C in a humid chamber, and then
examined for precipitin lines. Plates may be incubated at 2–8°C for another 24 hours to enhance the
precipitin lines.
An important consideration is the need for experienced personnel to interpret the AGID. Interpretation of
AGID results is dependent on the antigen used. Examples of AGIDs with different antigen preparations and
a guide for interpretation of the results can be found in Adams et al., 1983.
b)
Enzyme-linked immunosorbent assay (a prescribed test for international trade)
Currently, there are over 30 different ELISAs reported for the detection of anti-MVV or anti-CAEV antibodies
in the sera of sheep or goats, respectively (Deandres et al., 2005). Most of these ELISAs are indirect ELISAs
(I-ELISA) although there are three reported competitive ELISAs (C-ELISA) using monoclonal antibodies
(Frevereiro et al., 1999; Herrmann et al., 2003a; 2003b; Houwers & Schaake, 1987). Half of I-ELISAs use
purified whole virus preparations for antigen whereas the other half use recombinant protein and/or synthetic
peptide antigens. A few of the I-ELISAs have shown high sensitivity and specificity against a standard of
comparison, western blot analysis or radio-immunoprecipitation (Kwang et al., 1993; Rosati et al., 1994;
Saman et al., 1999). When compared with radio-immunoprecipitation, one C-ELISA has shown high
sensitivity and specificity both in sheep and goats in the USA suggesting that this one test can be used for
both MVV and CAEV US surveillance (Herrmann et al., 2003a; 2003b). Although ELISAs have been used for
several years in some European countries (Pépin et al., 1998) in control and eradication schemes of MVV in
sheep (Motha & Ralston, 1994) and CAEV in goats, AGID remains the most frequently used test.
Whole-virus antigen preparations are produced by differential centrifugation of supernatants from infected
cell cultures and detergent treatment of purified virus, and are coated on microplates (Dawson et al., 1982;
Simard & Briscoe, 1990a; Zanoni et al., 1994). Whole-virus preparations should contain both gp135 and
p28. Recombinant antigens or synthetic peptides are usually produced from whole or partial segments of the
gag or envelope genes and may be used in combination (Kwang et al., 1993; Power et al., 1995; Rosati et
al., 1994; Saman et al., 1999). Thus, recombinant gag or envelope gene products fused with glutathione Stransferase fusion protein antigen that have been produced in Escherichia coli provide a consistent source of
antigen for international distribution and standardisation.
The ELISA technique is also applicable to colostrum or milk, and some studies have evaluated paired serum
and milk samples. Because colostrum and milk are sources of CAEV transmission, the testing of milk
samples for anti-CAEV or anti-MVV antibody would not provide timely information for the prevention of
transmission, especially to offspring from the immediate gestation (Knowles, 1997).
The ELISA is performed at room temperature (~25°C) and is easy to perform in laboratories that have the
necessary equipment (microplate reader) and reagents. It is convenient for large-scale screening, as it is a
reliable and quantitative technique for demonstrating small ruminant lentiviruses (SRLVs) antibodies in
sheep and goats. It requires relatively pure antigen. One disadvantage of several ELISAs is that many have
not been validated against a standard of comparison such as western blot analysis or radioimmunoprecipitation. The OIE recommends the validation of 1000 negatives and 300 positives using a
standard of comparison such as western blot analysis or radio-immunoprecipitation, and to date, only one
ELISA has met these testing standards (Zanoni et al., 1994).
For I-ELISA, wells of the microplate are coated with antigen. Diluted serum samples are added to the wells
and react to antigens bound to the solid support. Unbound material is removed by washing after a suitable
incubation period. Conjugate (e.g. horseradish-peroxidase-labelled anti-ruminant Ig) reacts with specific
antibodies bound to the antigen. Unreacted conjugate is removed by washing after a suitable incubation
period. Enzyme substrate is added. The rate of conversion of substrate is proportional to the amount of
bound antibodies. The reaction is terminated after a suitable time and the amount of colour development is
measured spectrophotometrically. A disadvantage of the I-ELISA is that test sera typically need to be diluted
1/50 or greater in order to lower the number of false positives.
Specific MAbs have been used in a C-ELISA for SRLVs to capture gp135 or p28 as antigen (Frevereiro et
al., 1999; Herrmann et al., 2003a; 2003b; Houwers & Schaake, 1987; Ozyoruk et al., 2001). C-ELISA
overcomes the problem of antigen purity, as the specificity of this test depends on the MAb epitope. For CELISA, sample sera containing anti-SRLV antibodies inhibit binding of enzyme–labelled MAb to SRLV
antigen coated on the plastic wells. Binding of the enzyme-labelled MAb conjugate is detected by the
addition of enzyme substrate and quantified by subsequent colour product development. Strong colour
development indicates little or no blockage of enzyme-labelled MAb binding and therefore the absence of
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SRLV antibodies in sample sera. In contrast, weak colour development due to the inhibition of the enzymelabelled MAb binding to the antigen on the solid phase indicates the presence of SRLV antibodies in sample
sera. The format of the C-ELISA requires that serum antibodies must bind to or bind in close proximity to the
specific MAb epitope.
•
Materials and reagents
Microtitre plates with 96 flat-bottomed wells, freshly coated or previously coated with SRLV antigen;
microplate reader (equipped with 405, 450, 490 and 620 nm filters); 37°C humidified incubator; 1-, 8- and
12-channel pipettes with disposable plastic tips; microplate shaker (optional); fridge; freezer.
Positive and negative control sera; conjugate (e.g. ruminant anti-immunoglobulin labelled with peroxidase);
tenfold concentration of diluent (e.g. phosphate buffered saline/Tween); distilled water; 10× wash solution;
substrate or chromogen (e.g. ABTS [2,2’-azino-bis-(3-ethylbenzothiazoline-6-sulphonic acid)] or TMB
[3,3’,5,5’-tetramethylbenzidine]); stop solution (e.g. detergent, sulfuric acid).
•
Indirect ELISA: test procedure
i)
Dilute the serum samples, including control sera, to the appropriate dilution (e.g. 1/20) and distribute
0.1–0.2 ml per well (in duplicate if biphasic ELISA). Control sera are positive and negative sera
provided by the manufacturer and an internal positive reference serum from the laboratory in order to
compare the titres between different tests.
ii)
Cover the plate with a lid and incubate at room temperature or 37°C for 30–90 minutes. Empty the
contents and wash three times in washing solution at room temperature.
iii)
Add the appropriate dilution of freshly prepared conjugate to the wells (0.1 ml per well). Cover each
plate and incubate as in step ii. Wash again three times.
iv)
Add 0.1 ml of freshly prepared or ready-to-use chromogen substrate solution to each well (e.g. ABTS in
citrate phosphate buffer, pH 5.0, and 30% H2O2 solution [0.1 µl/ml]).
v)
Shake the plate; after incubation, stop the reaction by adding stopping solution to each well (e.g. 0.1 ml
sulphuric acid).
vi)
Read the absorbance of each well with the microplate reader at 405 nm (ABTS) or 450–620 nm (TMB).
The absorbance values will be used to calculate the results.
viii) Interpretation of the results
For commercial kits, interpretations and validation criteria are provided with the kit.
For example: calculate the mean absorbance (Ab) of the sample serum and of the positive (Abpos) and
negative (Abneg) control sera, and for each serum, calculate the percentage:
Ab - Abneg
x 100
Abpos - Abneg
Interpret the results as follows:
Ab <30%
negative serum
Ab 30–40%
doubtful serum
Ab >40%
positive serum
•
Competitive ELISA: test procedure
i)
Add 0.05 ml of undiluted serum and positive/negative controls to antigen-coated plate.
ii)
Incubate for 1 hour at room temperature.
iii)
Empty the plate and wash the plate three with diluted wash solution.
iv)
Add 0.05 ml of diluted antibody-peroxidase conjugate to each well. Mix well and incubate for
30 minutes at room temperature.
v)
After the 30-minute incubation, empty the plate and repeat the washing procedure described in step iii.
vi)
Add 0.05 ml of substrate solution (ex: TMB) to each well. Mix and cover plate with aluminium foil.
Incubate for 20 minutes at room temperature. Do not empty wells.
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vii)
Add 0.05 ml of stop solution to each well. Mix. Do not empty wells.
viii) Immediately after adding the stop solution, the plate should be read on a plate reader (620, 630 or
650 nm).
ix)
Interpretation of results
Example: Calculation: 100 – [(Sample Ab × 100)/(Mean negative control Ab)] = % inhibition.
For goats, if a test sample causes >33.2% inhibition, it is positive; if a test sample causes <33.2%
inhibition, it is negative. For sheep, if a test sample causes >20.9% inhibition, it is positive; if a test
sample causes <20.9% inhibition, it is negative.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
There are no biological products available. MAbs recognising conformational epitopes of the CAEV gp135 have
been described (Ozyoruk et al., 2001).
REFERENCES
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the p28 in immunodiffusion serology. Res. Vet. Sci., 40, 157–160.
ADAMS D.S., KLEVJER-ANDERSON P., CARLSON J.L., MCGUIRE T.C. & GORHAM J.R. (1983). Transmission and control
of caprine arthritis-encephalitis virus. Am. J. Vet. Res., 44, 1670–1675.
ALVAREZ V., ARRANZ J., DALTABUIT M., LEGINAGOIKOA I., JUSTE R.A., AMORENA B., DE ANDRES D., LUJAN L.L., BADIOLA
J.J. & BARRIATUA E. (2006). PCR detection of colostrum-associated Maedi-Visna virus (MVV) infection and
relationship with ELISA-antibody stutus in lambs. Res. Vet. Sci., 80, 226–234.
BANKS K.L., ADAMS D.S., MCGUIRE T.C. & CARLSON J. (1983). Experimental infection of sheep by caprine arthritisencephalitis virus and goats by progressive pneumonia virus. Am. J. Vet. Res., 44, 2307–2311.
CORK L.C. (1990). Pathology and epidemiology of lentiviral infection of goats. In: Maedi-Visna and Related
Diseases, Petursson G. & Hoff-Jørgensen R., eds. Kluwer Academic Press, Dordrecht, The Netherlands, 119–
127.
CRAWFORD T.B. & ADAMS D.S. (1981). Caprine arthritis-encephalitis: clinical features and presence of antibody in
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CRAWFORD T.B., ADAMS D.S., CHEEVERS W.P. & CORK L. C. (1980). Chronic arthritis in goats caused by a retrovirus.
Science, 207, 997–999.
CUTLIP R.C., JACKSON T.A. & LAIRD O.A. (1977). Immunodiffusion test for ovine progressive pneumonia. Am. J. Vet.
Res., 38, 1081–1084.
CUTLIP R.C., LEHMKUHL H.D., BROGDEN K.A. & MCCLURKIN A.W. (1986). Vasculitis associated with ovine
progressive pneumonia virus infection in sheep. Am. J. Vet. Res., 46, 61–64.
CUTLIP R.C., LEHMKUHL H.D., WOOD R.L. & BROGDEN K.A. (1985). Arthritis associated with ovine progressive
pneumonia. Am. J. Vet. Res., 46, 65–68.
DAWSON M. (1985). The detection of precipitating antibodies to lentivirus antigens in goat sera using two
immunodiffusion assays. In: Slow Viruses in Sheep, Goats and Cattle, Sharp J.M. & Hoff-Jørgensen R., eds.
Commission of the European Communities, EUR 8076, 233–238.
DAWSON M., BIRONT P. & HOUWERS D.J. (1982). Comparison of serological tests used in three state veterinary
laboratories to identify maedi-visna virus infection. Vet. Rec., 111, 432–434.
DEANDRES D., KLEIN D., WATT N.J., BERRIATUA E., TORSTEINSDOTTIR S., BLACKLAWS B.A. & HARKISS G.D. (2005).
Diagnostic tests for small ruminant lentiviruses. Vet. Microbiol., 107, 49–62.
FREVEREIRO M, BARROS S. & FUGULHA T. (1999). Development of a monoclonal antibody blocking-ELISA for
detection of antibodies against Maedi-Visna virus. J. Virol. Methods, 81, 101–108.
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HERRMANN L.M., CHEEVERS W.P., MCGUIRE T.C., ADAMS D.S., HUTTON M.M., GAVIN W.G. & KNOWLES D.P.A.
(2003a). A competitive-inhibition enzyme-linked immunosorbent assay (cELISA) for detection of serum antibodies
to caprine arthritis-encephalitis virus (CAEV): a diagnostic tool for successful eradication. Clin. Diagn. Lab.
Immunol., 10, 267–271.
HERRMANN L.M., CHEEVERS W.P., MARSHALL K.L., MCGUIRE T.C., HUTTON M.M., LEWIS G.S., KNOWLES D.P. (2003b).
Detection of serum antibodies to ovine progressive pneumonia virus in sheep by using a caprine arthritisencephalitis virus competitive-inhibition enzyme-linked immunosorbent assay. Clin. Diagn. Lab. Immunol., 10,
862–865.
HERRMANN L.M., HOTZEL I., CHEEVERS W.P., ON TOP K.P., LEWIS G.S. & KNOWLES D.P. (2004). Seven new ovine
progressive pneumonia virus (V) field isolates from Dubois Idaho sheep comprise part of OPPV clade II based on
surface envelope glycoprotein (SU) sequences. Virus Res., 102, 215–220.
HERRMANN L.M., MCGUIRE T.C., HOTZEL I., LEWIS G.S. & KNOWLES D.P. (2005). The surface envelope glycoprotein
(SU) is B-lymphocyte immunodominant in sheep naturally infected with ovine progressive pneumonia virus
(OPPV). Clin. Diagn. Lab. Immunol., 12, 797–800.
HERRMANN-HOESING L.M., WHITE S.N., LEWIS G.S., MOUSEL M.R. & KNOWLES D.P. (2007). Development and
validation of an ovine progressive pneumonia virus quantitative PCR. Clin. Vacc. Immunol., 14, 1274–1278.
HOUWERS D.J., GIELKENS A.L.J. & SCHAAKE J. (1982). An indirect enzyme-linked immunosorbent assay (ELISA) for
the detection of antibodies to maedi-visna virus. Vet. Microbiol., 7, 209.
HOUWERS D.J. & SCHAAKE J. (1987). An improved ELISA for the detection of antibodies to ovine and caprine
lentiviruses, employing monoclonal antibodies in a one-step assay. J. Immunol. Methods, 98, 151–154.
JOHNSON L.K., MEYER A.L. & ZINK M.C. (1992). Detection of ovine lentivirus in seronegative sheep by in situ
hybridization, PCR and cocultivation with susceptible cells. Clin. Immunol. Immunopathol., 65, 254–260.
KARR B.M., CHEBLOUNE Y., LEUNG K. & NARAYAN O. (1996). Genetic characterization of two penotypically distinct
North American ovine lentiviruses and their possible origin from caprine-arthritis encephalitis virus. Virology, 225,
1–10.
KNOWLES D.P. (1997). Laboratory diagnostic tests for retrovirus infections of small ruminants. Vet. Clin. North Am.
Food Anim. Pract., 13, 1–11.
KNOWLES D.P., EVERMANN J.F., SCHROPSHIRE C., VANDER SCHALIE J., BRADWAY D., GEZON H.M. & CHEEVER W.P.
(1994). Evaluation of agar gel immunodiffusion serology using caprine and ovine lentiviral antigens for detection
of antibody to caprine-arthritis encephalitis virus. J. Clin. Microbiol. 32, 243–245.
KNOWLES D., CHEEVERS W., MCGUIRE T., STEM T. & GORHAM J. (1990). Severity of arthritis is predicted by antibody
response to gp135 in chronic infection with caprine arthritis-encephalitis virus. J. Virol., 64, 2396–2398.
KWANG J., KEEN J., CUTLIP R.C. & LITTLEDIKE E.T. (1993). Evaluation of an ELISA for detection of ovine progressive
pneumonia antibodies using a recombinant transmembrane envelope protein. J. Vet. Diagn. Invest., 5, 189–193.
LEROUX C., CHASTANG J., GREENLAND T. & MORNEX J.F. (1997). Genomic heterogenetity of small ruminant
lentiviruses: existence of heterogeneous populations in sheep and of the same lentiviral genotypes in sheep and
goats. Arch. Virol., 142, 1125–1137.
MOTHA M.J. & RALSTON J.C. (1994). Evaluation of ELISA for detection of antibodies to CAE in milk. Vet. Microbiol.,
38, 359–367.
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pathologic and virologic studies on the naturally occurring disease. Am. J. Vet. Res., 42, 1554–1559.
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intrathoracic, intracerebral and intra-articular infections. Am. J. Vet. Res., 42, 1560–1564.
OZYORUK F., CHEEVERS W.P., HULLINGER G.A., MCGUIRE T.C., HUTTON M. & KNOWLES D.P. (2001). Monoclonal
antibodies to conformational epitopes of the surface glycoprotein of caprine arthritis-encephalitis virus: potential
application to competitive-inhibition enzyme-linked immunosorbent assay for detecting antibodies in goat sera.
Clin. Diagn. Lab. Immunol., 8, 44–51.
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PÉPIN M., VITU C., RUSSO P., MORNEX J.F. & PETERHANS E. (1998). Maedi-visna virus infection in sheep: a review.
Vet. Res., 29, 341–367.
PETERHANS E., GREENLAND T., BADIOLA J., HARKISS G., BERTONI G., AMORENA B., ELIASZEWICZ M., JUSTE R., KRASSNIG
R., LAFONT J.P., LENIHAN P., PETURSSON G., PRITCHARD G., THORLEY G., VITU C., MORNEX J.F. & PÉPIN M. (2004).
Routes of transmission and consequences of small ruminant lentiviruses (SRLVs) infection and eradication
schemes. Vet. Res., 35, 257–274.
POWER C., RICHARDSON S., BRISCOE M. & PASICK J. (1995). Evaluation of two recombinant Maedi-Visna virus
proteins for use in an enzyme-linked immunosorbent assay for the detection of serum antibodies to ovine
lentiviruses. Clin. Diagn. Lab. Immunol., 2, 631–633.
RIMSTAD E., EAST N., DEROCK E., HIGGINS J. & PEDERSEN N.C. (1994). Detection of antibodies to caprine
arthritis/encephalitis virus using recombinant gag proteins. Arch. Virol., 134, 345–356.
ROLAND M., MOONEY J., VALAS S., PERRIN G. & MAMOUN R.Z. (2002). Characterization of an Irish caprine lentivirus
strain-SRLV phylogeny revisited. Virus Res., 85, 29–39.
ROSATI S., KWANG J., TOLARI F. & KEEN J.E. (1994). A comparison of whole virus and recombinant transmembrane
ELISA and immunodiffusion for detection of ovine lentivirus antibodies in Italian sheep flocks. Vet. Res. Commun.,
18, 73–80.
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BADIOLA J.J. (1999). A new sensitive serological assay for detection of lentivirus infections in small ruminants, Clin.
Diagn. Lab. Immunol., 6, 734–740.
SHAH C., BÖNI J., HUDER J.B., VOGT H.R., MÜLHERR J., ZANONI R., MISEREZ R., LUTZ H. & SCHÜPBACH J. (2004a).
Phylogenetic analysis and reclassification of caprine and ovine lentiviruses based on 104 new isolates: evidence
for regular sheep-to-goat transmission and worldwide propagation through lifestock trade. Virology, 319, 12–26.
SHAH C., HUDER J.B., BÖNI J., SCHÖNMANN M., MÜHLHERR J., LUTZ H. & SCHÜPBACH J. (2004b). Direct evidence for
natural transmission of small ruminant lentiviruses of subtype A4 from goats to sheep and vice versa. J. Virol., 78,
7518–7522.
SIMARD C.L. & BRISCOE M.R. (1990a). An enzyme-linked immunosorbent assay for detection of antibodies to
maedi-visna virus in sheep. A simple technique for production of antigen using sodium dodecyl sulfate treatment.
Can. J. Vet. Res., 54, 446–450.
SIMARD C.L. & BRISCOE M.R. (1990b). An enzyme-linked immunosorbent assay for detection of antibodies to
Maedi-visna virus in sheep. Comparison to conventional agar gel immunodiffusion test. Can. J. Vet. Res., 54,
451–456.
TERPSTRA C. & DE BOER G.F. (1973). Precipitating antibodies against maedi-visna virus in experimentally infected
sheep. Archiv für die gesamte Virusforschung, 43, 53–62.
VALAS S., BENOIT C., GUIONAUD C., PERRIN G. & MAMOUN R.Z. (1997). North American and French caprine arthritisencephalitis viruses emerge from ovine maedi-visna viruses. Virology, 237, 307–318.
ZANONI R.G. (1998). Phylogenetic analysis of small ruminant lentiviruses. J. Gen. Virol., 79, 1951–1961.
ZANONI R.G., VOGT H.R., POHL B., BOTTCHER J., BOMMELI W. & PETERHANS E. (1994). An ELISA based on whole
virus for the detection of antibodies to small-ruminant lentiviruses. J. Vet. Med. B, 41, 662–669.
*
* *
NB: There are OIE Reference Laboratories for Caprine arthritis/encephalitis & Maedi-visna
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for caprine arthritis/encephalitis & Maedi-visna
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.7.5.
CONTAGIOUS AGALACTIA
SUMMARY
Contagious agalactia is a serious disease syndrome of sheep and goats that is characterised by
mastitis, arthritis, keratoconjunctivitis and, occasionally, abortion. Mycoplasma agalactiae is the
main cause of the disease in sheep and goats, but M. capricolum subsp. capricolum (Mcc),
M. mycoides subsp. capri (Mmc) (formerly named M. mycoides subsp, mycoides LC [LC = large
colonies]) and M. putrefaciens produce a clinically similar disease, more often in goats, which may
be accompanied by pneumonia. Antibodies to Mmc and Mcc have been detected in South
American camelids (alpacas, llamas and vicunas), but no mycoplasmas have yet been isolated.
Identification of the agent: Definitive diagnosis requires the isolation of the causative
mycoplasmas from the affected animals, which are then identified by biochemical, serological and,
increasingly, molecular tests such as the polymerase chain reaction. Samples of choice include
milk, conjunctival and ear swabs, and joint fluid. All four mycoplasmas grow relatively well in most
mycoplasma media although M. agalactiae shows a preference for organic acids such as pyruvate
as substrates.
Serological tests: Detection of antibodies in serum by complement fixation test or enzyme-linked
immunosorbent assay (ELISA) provides rapid diagnosis of disease, but may not be very sensitive in
chronically affected herds and flocks. Indirect ELISAs have been used routinely in control
programmes for screening herds for M. agalactiae. Confirmation of infection by isolation and
identification is usually necessary in areas believed to be free of contagious agalactia. Serological
tests are not widely available for M. putrefaciens.
Requirements for vaccines and diagnostic biologicals: Commercial vaccines for M. agalactiae,
inactivated with formalin, are widely used in southern Europe, but are not considered to be very
efficacious. Under experimental conditions, M. agalactiae vaccines inactivated with saponin or
phenol have been shown to be more protective than formalised preparations. Live vaccines for
M. agalactiae are used in Turkey, where they are reported to be more protective than inactivated
vaccines. A commercial vaccine containing M. agalactiae, Mmc and Mcc is available. Autogenous
vaccines for Mmc and, occasionally, for Mcc are believed to be used in some countries. No
vaccines exist for M. putrefaciens, as the disease it causes is not considered to be sufficiently
serious or widespread.
A. INTRODUCTION
Contagious agalactia is a disease of sheep and goats that is characterised by mastitis, arthritis and
keratoconjunctivitis, and has been known for nearly 200 years. It occurs in Europe, western Asia, the United
States of America (USA) and North Africa, and is mainly caused by Mycoplasma agalactiae (Bergonier et al.,
1997). In recent years, M. capricolum subsp. capricolum (Mcc) and M. mycoides subsp. capri1 (formerly
M. mycoides subsp. mycoides LC [LC = large colonies]) have also been isolated in many countries from sheep
and goats with mastitis and arthritis. The clinical signs of these infections are sufficiently similar to be considered
indistinguishable from contagious agalactia. In addition, M. putrefaciens also causes mastitis and arthritis in
goats, which is very similar to that caused by M. agalactiae, Mmc and Mcc (Rodriguez et al., 1994). Furthermore,
the consensus of the working group on contagious agalactia of the EC COST2 Action 826 on ruminant
1
2
International Committee on Systematics of Prokaryotes Subcommittee on the Taxonomy of Mollicutes has proposed the
merging of these two subspecies into the single subsp: M. mycoides subsp. capri; a decision is pending.
European Cooperation in the field of Scientific and Technical Research.
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Chapter 2.7.5. — Contagious agalactia
mycoplasmoses, which met in Toulouse, France, in 1999, was that all four mycoplasmas should be considered as
causal agents of contagious agalactia.
Clinically, the disease caused by M. agalactiae is recognised by elevated temperature, inappetence and alteration
in the consistency of the milk in lactating ewes with decline and subsequent failure of milk production, often within
2–3 days, as a result of interstitial mastitis (Bergonier et al., 1997); lameness and keratoconjunctivitis affects
about 5–10% of infected animals. Fever is common in acute cases and may be accompanied by nervous signs,
but both signs are rare in the more frequently observed subacute and chronic infections. Pregnant animals may
abort. Mycoplasma agalactiae may occasionally be found in lung lesions (Loria et al., 1999), but pneumonia is not
a consistent finding. Bacteraemia is common, particularly for Mmc and Mcc and could account for the isolation of
the organism from sites where it is only transiently present.
Mastitis, arthritis, pleurisy, pneumonia, and keratoconjunctivitis may all result from infection with Mmc, which has
one of the widest geographical distribution of ruminant mycoplasmas, being found on all continents where small
ruminants are kept and wherever contagious agalactia and caprine pleuropneumonia are reported (Da Massa et
al., 1983; Nicholas, 2002); however the lack of diagnostic facilities for mycoplasma diseases in many countries
means that it is probably under reported. Mmc is mostly confined to goats but has occasionally been isolated from
sheep with reproductive disease and cattle with arthritis or respiratory disease. Cases usually occur sporadically,
but the disease may persist and spread slowly within a herd. After parturition, the opportunity for spread in milking
animals increases, and kids ingesting infected colostrum and milk become infected. The resulting septicaemia,
with arthritis and pneumonia, causes high mortality in kids (Bergonier et al., 1997; Da Massa et al., 1983).
Mcc is widely distributed and highly pathogenic, particularly in North Africa but the frequency of occurrence is low
(Bergonier et al., 1997). Goats are more commonly affected than sheep, and clinical signs of fever, septicaemia,
mastitis, and severe arthritis may be followed rapidly by death (Bergonier et al., 1997; Bolske et al., 1988).
Pneumonia may be seen at necropsy. The severe joint lesions seen in experimental infections with this organism
are accompanied by intense periarticular subcutaneous oedema affecting tissues some distance from the joint
(Bolske et al., 1988).
Mycoplasma putrefaciens is common in milking goat herds in western France where it can be isolated from
animals with and without clinical signs (Mercier et al., 2001). It has also been associated with a large outbreak of
mastitis and agalactia leading to severe arthritis in goats accompanied by abortion and death without pyrexia in
California, USA (Bergonier et al., 1997). Mycoplasma putrefaciens was the major finding in an outbreak of
polyarthritis in kids in Spain (Rodriguez et al., 1994).
Antibodies to Mmc and Mcc, but not M. agalactiae, have been detected in South American camelids, including
llamas, alpacas and vicunas, but as yet no mycoplasmas have been isolated (Nicholas, 1998). These camelids
are affected by a range of mycoplasma-like diseases, including polyarthritis and pneumonia, so it is likely that
mycoplasmas including Mmc and Mcc may be found in the future.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agents
a)
Selection of samples
Preferred samples from living animals include: nasal swabs and secretions; milk from mastitic females or
from apparently healthy females where there is a high rate of mortality/morbidity in kids; joint fluid from
arthritic cases; conjunctival swabs from cases of ocular disease; and blood for antibody detection from
affected and non-affected animals (Nicholas & Baker, 1998). The ear canal has also been shown to be a
source of pathogenic mycoplasmas, although in practice the presence of nonpathogenic mycoplasmas at
this site may make confirmation difficult (Nicholas & Baker, 1998). Mycoplasmas may be isolated from the
blood during the acute stage of the disease when there is mycoplasmaemia. From dead animals, samples
should include: udder and associated lymph nodes, joint fluid, lung tissue (at the interface between diseased
and healthy tissue) and pleural/pericardial fluid. Samples should be dispatched quickly to a diagnostic
laboratory in a moist and cool condition. All four causative mycoplasmas are relatively easy to isolate from
internal organs, joints and milk and grow well in most mycoplasma media, producing medium to large
colonies in 3–4 days.
b)
Mycoplasma medium
The usual techniques used in the isolation of mycoplasmas apply to all four causative organisms (Nicholas &
Baker, 1998). Many media have been reported to grow the causative mycoplasmas. Improved growth rates
of M. agalactiae have been seen in media containing organic acids such as pyruvate and isopropanol (Khan
et al., 2004). The formulation of PRM medium (Khan et al., 2004) is as follows:
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Chapter 2.7.5. — Contagious agalactia
Heat inactivated porcine serum 100 ml/litre, special peptone 20 g/litre, yeast extract 5 g/litre, glycerol
5 g/litre, sodium chloride 5 g/litre, HEPES 9 g/litre, fresh yeast extract 100 ml/litre, sodium pyruvate 5 g/litre,
12.5 ml of 0.2% phenol red and ampicillin (200,000 International Units/ml. Make up to 1 litre in distilled water
and sterilise by filtration. Adjust the pH of the broth medium to 7.6. Prepare solid medium by adding 10 g of
LabM agar No. 1 (Bury UK, or agar of equivalent quality) and dispense into sterile Petri dishes.
Thallium acetate (250 mg/litre), which is toxic and inhibitory to some mycoplasmas but not those causing
contagious agalactia, may be a necessary component of the transport medium to reduce bacterial
contamination from clinical samples, but should be omitted once the mycoplasmas begin to grow in vitro. A
satisfactory alternative to thallium acetate may be colistine sulphate (37.5 mg/litre).

Test procedure
i)
Make tenfold dilutions (101–106) of the liquid sample (milk, synovial fluid, conjunctival and ear swabs)
or tissue homogenate in appropriate broth medium.
ii)
Spread a few drops of each sample on the agar medium and dispense a 10% (v/v) inoculum into broth
medium.
iii)
Streak swabs directly on to agar medium.
iv)
Incubate inoculated broths (optimally with gentle shaking) and agar media at 37°C in humidified
atmosphere with 5% carbon dioxide.
v)
Examine broths daily for signs of growth (indicated by a fine cloudiness or opalescence) or changes in
pH indicated by a colour change and examine agar media under × 35 magnification for typical ‘fried
egg’ colonies.
vi)
If no mycoplasma growth is seen after 7 days, subculture a 10% (v/v) inoculum of broth into fresh broth
and spread about 50 µl of this on to agar media.
vii)
Repeat as for step v. If no mycoplasmas are seen after 21 days’ incubation, consider the results to be
negative.
viii) If bacterial contamination results (seen as excessive turbidity), filter sterilise by passing 1 ml of
contaminated broth through a 0.45 µm filter into fresh broth medium.
Clinical samples frequently contain more than one mycoplasma species so clone purification of colonies is
often considered necessary before performing biochemical and serological identification, in particular the
growth and film inhibition tests (GIT and FIT, respectively). However, cloning is a lengthy procedure taking at
least 2 weeks. The immunofluorescence test (Bradbury, 1998), dot immunobinding tests (Poumarat, 1998)
and, more recently, polymerase chain reaction (PCR) tests (see Section B.1.e) do not require cloning as
these tests can detect the pathogenic mycoplasmas in mixed cultures, saving a great deal of time.
c)
Biochemical tests
The first test that should be performed on the cloned isolates is sensitivity to digitonin, which separates
mycoplasmas from acholeplasmas; the latter are ubiquitous contaminants that can overgrow the
mycoplasmas of interest. Growth in liquid medium containing glucose (1%), arginine (0.2%), and
phenolphthalein diphosphate (0.01%), on solid medium containing horse serum or egg yolk for the
demonstration of film and spots, and on casein agar or coagulated serum agar to test for proteolysis, are
among the most useful tests for differentiating the four mycoplasmas (Poveda, 1998). These biochemical
characteristics, however have been increasingly found to be variable for the individual mycoplasmas and
have little diagnostic value. The most impressive biochemical characteristic that differentiates
M. putrefaciens from all other mycoplasmas is the odour of putrefaction it produces in broth culture. Other
features that may be helpful include: film and spot production seen on the surface of the broth and solid
media caused by M. agalactiae and to a lesser extent by M. putrefaciens; and the proteolytic activity of Mcc
and MmmLC on casein and coagulated serum.
A rapid and highly convenient biochemical test that exploits the C8-esterase activity of M. agalactiae has
been reported (Khan et al., 2001). The mycoplasma forms red colonies on agar media within 1 hour of
adding the chromogenic substrate, SLPA-octanoate (a newly synthesised ester formed from a C8 fatty acid
and a phenolic chromophore). This activity is shared with M. bovis, although this mycoplasma is rarely found
in small ruminants. Isolates need not be cloned as M. agalactiae can be detected easily in mixed cultures. If
necessary PCRs can be used to distinguish rapidly M. agalactiae from M. bovis (see Section B.1.e).
d)
Serological identification
Identification of isolates using specific antisera is usually carried out with the GIT, FIT (Poveda & Nicholas,
1998) or the indirect fluorescent antibody (IFA) test (Bradbury, 1998). A recently developed dot
immunobinding test, which is carried out in microtitre plates, offers many improvements over the other
serological tests such as rapidity and higher throughputs (Poumarat, 1998) but requires subjective
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Chapter 2.7.5. — Contagious agalactia
judgements of staining intensity. For M. agalactiae, film inhibition may often be more reliable as growth
inhibition is not seen with all isolates; it can also be used for serodiagnosis. Film production by the
mycoplasma may be enhanced by the incorporation of 10% egg yolk suspension into the solid medium.

Test procedure
i)
Inoculate at least two dilutions of 48-hour cloned broth cultures (10–1 and 10–2) on to predried agar
media by allowing 50 µl of the cultures to run down the tilted plates using the ‘running drop’ technique
(Poveda & Nicholas, 1998). Remove any excess liquid with a pipette.
ii)
Allow the plates to dry. It is possible to apply two or three well separated running drops to each 90 mm
diameter plate.
iii)
Apply predried filter paper discs containing 30 µl of specific antiserum to the culture; ensure good
separation of discs (at least 30 mm).
iv)
Incubate the plates as for mycoplasma culture and examine daily by eye against a light background.

Interpretation of the results
A zone of inhibition over 2 mm, measured from the paper disc to the edge of mycoplasma growth is
considered to be significant. Partial inhibition can occur with weak antiserum or where there are mixed
cultures. Stronger reactions can be obtained if about 60 µl of antisera is added to 6 mm diameter wells made
in the agar with a cork borer or similar device (Poveda & Nicholas, 1998).
In the IFA test, specific antisera are applied to colonies on solid medium. Homologous antiserum remains
attached after washing and is demonstrated by adding fluorescein-conjugated antiglobulin, washing, and
viewing the colonies with an epifluorescence microscope (Bradbury, 1998). Controls should include known
positive and known negative control organisms, and a negative control serum. However like the
immunobinding tests subjective judgements are required to assess staining intensity.
Antisera for these serological tests have traditionally been prepared against the type strains of the various
Mycoplasma species, and most field isolates have been readily identified using these antisera. As more
strains have been examined, however, some have been found to react poorly with these antisera, while
reacting well with antisera to other representative strains of the species. Intraspecies variation in antigenic
composition has not been reported for M. putrefaciens, but occurs to some degree with M. agalactiae and
with Mcc strains. Thus, diagnostic laboratories may need to have several antisera to enable all strains of the
species to be identified.
e)
Nucleic acid recognition methods
PCR assays are routinely used in many laboratories and are extremely sensitive. They can provide a rapid
early warning system when carried out on clinical samples, enabling a full investigation to take place when
results are positive. However negative results should not be considered definitive. Several PCRs specific for
M. agalactiae have been developed and show similar levels of sensitivity, although they are based on
different gene sequences (Bashiruddin et al., 2005; Dedieu et al., 1995; Subrahamaniam et al., 1998; Tola et
al., 1997a). They can be used directly on nasal, conjunctival, synovial and tissue samples; they have been
used on milk samples where they have been reported to be more sensitive than culture (Tola et al., 1997a),
although occasionally undefined inhibitors may interfere with the test. PCRs can also be used, more reliably,
on mycoplasmas growing in culture; a 24 hour enrichment of the mycoplasma in the appropriate medium
greatly facilitates PCR detection even in the presence of bacterial contamination (Nicholas, 2002). A newly
described PCR based method called denaturing gradient gel electrophoresis (DGGE) that uses
mycoplasma-specific primers is capable of identifying the majority of small ruminant mycoplasmas including
all the causative agents of contagious agalactia by their migration pattern (McAuliffe et al., 2005). A positive
PCR result, particularly in an area previously free of contagious agalactia, should be confirmed by isolation
and identification of the mycoplasma using standard procedures.
Individual PCRs have been reported for Mmc and Mcc (Bashiruddin et al., 1994) and M. putrefaciens
(Peyraud et al., 2003) respectively. In addition a multiplex test has been described which can detect
simultaneously M. agalactiae, Mcc and Mmc (Greco et al., 2001).

Test procedure
The following primers based on the uvrC gene have been shown to be specific for M. agalactiae
(Subrahamanian et al., 1998). PCRs may need to be optimised in each laboratory. Positive and negative
control DNA should be run in each assay.
MAGAUVRC1-L
MAGAUVRC1-R
990
CTC-AAA-AAT-ACA-TCA-ACA-AGC
CTT-CAA-CTG-ATG-CAT-CAT-AA
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Chapter 2.7.5. — Contagious agalactia
i)
Extract DNA from Mycoplasma isolates or clinical material using the appropriate method (Bolske et al.,
1988).
ii)
Carry out PCR methods in 50 µl reaction mixtures containing: 1 µl of sample DNA, 20 pmol of each
primer (see above), 1 mM each dNTP, 10 mM Tris/HCl, pH 8.3, 1.5 mM MgCl2, 50 mM KCl and
1.25 mM U Taq DNA polymerase.
iii)
Subject the mixture to 35 amplification cycles in a thermal cycler with the following parameters:
30 seconds at 94°C, 30 seconds at 50°C annealing temperature and 1 minute at 72°C.
iv)
Analyse the PCR products by electrophoresis on a 0.7% agarose at 110 V for 2 hours and visualise by
staining with ethidium bromide. A 1.7 kb fragment indicates the presence of M. agalactiae.
2.
Serological tests
a)
Complement fixation
A standard complement fixation test (CFT) for M. agalactiae has also been applied to other mycoplasmas
involved in the contagious agalactia syndrome (Bergonier et al., 1997). Antigens are prepared from washed
organisms, standardised by opacity, and lysed, either ultrasonically or by using sodium lauryl sulphate followed
by dialysis. Sera are inactivated at 60°C for 1 hour, and the test is carried out in microtitre plates with overnight
fixation in the cold or at 37°C for 3 hours. The haemolytic system is added, and the test is read after complete
lysis is shown by the antigen control. A positive result is complete fixation at a serum dilution of 1/40 or greater
for the following mycoplasmas: M. agalactiae, Mcc, and Mmc. The CFT is regarded as a herd test and at least
ten sera are tested from each herd, preferably from acute and convalescent cases.
Some sera from healthy flocks react in the CFT using M. agalactiae up to a serum dilution of 1/20, but rarely
react with the other two antigens. However, in flocks infected with M. agalactiae, sera giving a homologous
reaction at 1/80 may cross-react at up to 1/40, the positive threshold, with the other two antigens. It is often
difficult to perform the CFT if the quality of the test sera is poor; where possible, the enzyme-linked
immunosorbent assay (ELISA) is preferred.
b)
Enzyme-linked immunosorbent assay
ELISAs using sonicated or Tween-20-treated antigens have been reported to be more sensitive than the
CFT for the detection of antibody to M. agalactiae (Bergonier et al., 1997). Problems of nonspecificity have
been overcome by the use of monoclonal or protein G conjugates in the ELISA (Lambert et al., 1998). The
use of these conjugates enables the testing of sera from a wide range of mammalian species, including
camelids. A number of commercial ELISA kits are now available and these are being used for large-scale
surveys in France and the United Kingdom (Bergonier et al., 1997; Nicholas, 1998). In a ring trial of
serological tests for M. agalactiae organised in 1998 under the auspices of the EC COST Action 826 on
ruminant mycoplasmoses, commercial ELISAs performed better than ‘home-made’ kits.
ELISAs are not widely available for the other three causative mycoplasmas although ‘home-made’ assays
are carried out by some laboratories.
c)
Immunoblotting test
Immunoblotting tests have been reported as the most sensitive and specific test for M. mycoides subsp.
mycoides SC, the cause of contagious bovine pleuropneumonia (see chapter 2.4.9). Immunoblotting tests
have also been described for M. agalactiae (Nicholas, 1998; Tola et al., 1997b). Strong bands at
approximately 80 and 55 kDa were seen with sera with antibodies to M. agalactiae, while sera from healthy
flocks show no bands or very faint bands of different sizes. Diluting the sera to 1/50 improves the
discrimination between positive and negative sera (Nicholas, 1998).
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
Vaccines for the prevention of contagious agalactia due to M. agalactiae are used widely in the Mediterranean
countries of Europe and in western Asia. No single vaccine has been universally adopted, and no standard
methods of preparation and evaluation have been applied.
1.
Vaccines for Mycoplasma agalactiae infection
a)
Inactivated vaccines for Mycoplasma agalactiae infection
In Europe, where live vaccines for M. agalactiae are not acceptable, attention has focused on the use of
killed organisms, mostly using formalin and an adjuvant such as aluminium hydroxide in an oil emulsion. The
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Chapter 2.7.5. — Contagious agalactia
titres of the preparations, before inactivation, are very high (108–1010 colony-forming units per ml) and are
derived from laboratory strains. Some products are available commercially including a trivalent preparation
containing M. agalactiae, Mcc and Mmc but there are few data on their efficacy. A formalin-inactivated oil
emulsion vaccine was shown to be immunogenic and protective in a small trial in lactating sheep and also
prevented transmission of M. agalactiae (Greco et al., 2002).
It is possible that in some instances the apparent lack of protection given by vaccines could be the result of
animals being infected with one of the other four mycoplasmas involved in the contagious agalactia
syndrome (Gil et al., 1999). A multivalent formalin inactivated vaccine incorporating all four causative
mycoplasmas and adjuvanted with saponin and aluminium hydroxide appears beneficial in preliminary trials
(Ramirez et al., 2001).
More recently vaccines inactivated with phenol or with saponin have given superior protection against
experimental infections compared with formalin, sodium hypochlorite or heat-inactivated vaccines (Tola et
al., 1999).
b)
Live attenuated vaccines for Mycoplasma agalactiae infection
Live attenuated vaccines against M. agalactiae have been used in Turkey for many years and have been
reported to provide better protection in ewes and their lambs than inactivated vaccines (Nicholas, 2002).
However they can produce a transient infection with shedding of mycoplasma. Live vaccines should not be
used in lactating animals and should be part of a regional plan in which all flocks from which animals are
likely to come into contact be vaccinated at the same time.
2.
Vaccines for Mycoplasma mycoides subsp. capri infection
There is little recent published information on the availability of vaccines for Mmc although it is believed that
inactivated vaccines are widely used in many Mediterranean countries and in Asia suggesting that their
production and use is localised (Bergonier et al., 1997). Saponised vaccines have been reported in India which
provoke a strong antibody response and show some protection (Sunder et al., 2002).
3.
Mycoplasma capricolum subsp. capricolum and M. putrefaciens
Although infections with Mcc and M. putrefaciens can be severe, their prevalence is relatively low and, as might
be expected, little or no work appears to have been carried out on preventive vaccination for these infections.
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AYLING R.D., NICHOLAS R.A.J., THIAUCOURT F. & SACHSE K. (2005). Evaluation of PCR systems for the identification
and differentiation of Mycoplasma agalactiae and Mycoplasma bovis: A collaborative trial. Vet. J., 169, 268–275.
BASHIRUDDIN J.B., TAYLOR T.K. & GOULD A.R. (1994). A PCR-based test for the specific identification of
Mycoplasma mycoides subsp. mycoides SC in clinical material. J. Vet. Diagn. Invest., 6, 428–434.
BERGONIER D., BERTHOLET X. & POUMARAT F. (1997). Contagious agalactia of small ruminants: current knowledge
concerning epidemiology, diagnosis and control. Rev. sci. tech. Off. int. Epiz., 16, 848–873.
BOLSKE G., MSAMI H., HUMLESLO N.E, ERNO H. & JOHNSSON L. (1988). Mycoplasma capricolum in an outbreak of
polyarthritis and pneumonia in goats. Acta Vet. Scand., 29, 331–338.
BRADBURY J.M. (1998). Identification of mycoplasmas by immunofluorescence. In: Mycoplasma Protocols, Miles
R.J. & Nicholas R.A.J., eds. Humana Press, Totowa, USA, 119–125.
DA MASSA A.J., BROOKS D.L. & ADLER H.E. (1983). Caprine mycoplasmosis: widespread infection in goats with
Mycoplasma mycoides subsp. mycoides (large-colony type). Am. J. Vet. Res., 44, 322–325.
DEDIEU L., MADY V. & LEFEVRE P. C. (1995). Development of two PCRs for the identification of mycoplasmas
causing contagious agalactia. FEMS Microbiol. Lett., 129, 243–250.
GRECO G., CORRENTE M., BUONOVOGLIA D., ALIBERTI A. & FASANELLA A. (2002). Inactivated vaccine induces
protection against Mycoplasma agalactiae infection in sheep. Microbiologica, 25, 17–20.
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GRECO G., CORRENTE M., MARTELLA V., PRATELLI A. & BOUNOVOGLIA D. (2001). A mulitiplex PCR for the diagnosis of
contagious agalactia of sheep and goats. Mol. Cell. Probes, 15, 21–25.
GIL M.C., HERMOSA DE MENDOZA M., REY J., ALONSO J.M. POVEDA J.B. & HERMOSA DE MENDOZA J. (1999). Aetiology
of caprine contagious agalactia syndrome in Extramudura, Spain. Vet. Rec., 144, 24–25.
KHAN L., LORIA G., ABU-AMERO K., NICHOLAS R.A.J., HALABLAB M. & MILES R.J. (2001). Distinctive biochemical
characteristics of Mycoplasma agalactiae and Mycoplasma bovis. In: Mycoplasmas of Ruminants: Pathogenicity,
Diagnostics, Epidemiology and Molecular Genetics, Vol. 5, Poveda J.B., Fernandez A., Frey J. & Johansson K.E., eds. European Commission, Brussels, Belgium, 60–63.
KHAN L.A., LORIA G.R., RAMIREZ A.S., NICHOLAS R.A.J., MILES R.J. & FIELDER M.D. (2004). Biochemical
characterisation of some non fermenting, non arginine hydrolysing mycoplasmas of ruminants. Vet. Microbiol.,
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LAMBERT M., CALAMEL M., DU FOUR P., CABASSE E., VITU C. & PEPIN M. (1998). Detection of false-positive sera in
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LORIA G.R., SAMMARTINO C., NICHOLAS R.A.J &. AYLING R.D. (1999). In vitro susceptibility of field isolates of
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NICHOLAS R.A.J. (2002). Improvements in the diagnosis and control of diseases of small ruminants caused by
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detection of Mycoplasma putrefaciens, one of the agents of the contagious agalactia syndrome of goats. Mol.
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POUMARAT F. (1998). Identification of mycoplasmas by dot immunobinding on membrane filtration (MF Dot). In:
Mycoplasma Protocols, Miles R.J. & Nicholas R.A.J., eds. Humana Press, Totowa, USA, 113–118.
POVEDA J.B. (1998). Biochemical characteristics in mycoplasma identification. In: Mycoplasma Protocols, Miles
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RAMIREZ A S., DE LA FE C., ASSUNCAO P., GONZALEZ M. & POVEDA J.B. (2001). Preparation and evaluation of an
inactivated polyvalent vaccine against Mycoplasma spp on infected goats. In: Mycoplasmas of Ruminants:
Pathogenicity, Diagnostics, Epidemiology and Molecular Genetics, Vol. 5, Poveda J.B., Fernandez A., Frey J. &
Johansson K.-E., eds. European Commission, Brussels, Belgium, 154–157.
RODRIGUEZ J.L., POVEDA J.B., GUTIERREZ C., ACOSTA B. & FERNANDEZ A. (1994). Polyarthritis in kids associated with
Mycoplasma putrefaciens. Vet. Rec., 135, 406–407.
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SUBRAHAMANIAM S., BERGONIER D., POUMARAT F., CAPUAL S., SCHLATTER Y., NICOLET J. & FREY J. (1998). Species
identification of Mycoplasma bovis and Mycoplasma agalactiae based on the uvrC gene by PCR. Mol. Cell.
Probes, 12, 161–169.
SUNDER J., SRIVASTAVA N.C. & SINGH V.P. (2002) Preliminary trials on development of vaccine against Mycoplasma
mycoides subsp. mycoides type LC infection in goats. J. Appl. Anim. Res., 21, 75–80.
TOLA S., ANGIOI A., ROCCHIGIANI A.M., IDINI G., MANUNTA D., GALLERI G. & LEORI G. (1997a). Detection of
Mycoplasma agalactiae in sheep milk samples by polymerase chain reaction. Vet. Microbiol., 54, 17–22.
TOLA S., MANUNTA D., COCCO M., TURRININ F., ROCCHIGIANI A.M., IDINI G., ANGIOI A. & LEORI G. (1997b).
Characterisation of membrane surface proteins of Mycoplasma agalactiae during natural infection. FEMS
Microbiol. Lett., 154, 355–362.
TOLA S., MANUNTA D., ROCCA S., ROCCHIGIANI A.M., IDINI G., ANGIOI A. & LEORI G. (1999). Experimental vaccination
of against Mycoplasma agalactiae using different inactivated vaccine. Vaccine, 17, 2764–2768.
*
* *
NB: There is an OIE Reference Laboratory for contagious agalactia
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for contagious agalactia
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.7.6.
CONTAGIOUS CAPRINE PLEUROPNEUMONIA
SUMMARY
Contagious caprine pleuropneumonia (CCPP) is a severe disease of goats caused by Mycoplasma
capricolum subspecies capripneumoniae (Mccp). This organism is closely related to three other
mycoplasmas: M. mycoides subsp. mycoides large colonies (LC), M. mycoides subsp. capri, and
M. capricolum subsp. capricolum. Unlike true CCPP, which is confined to the thoracic cavity, the
disease caused by the latter three mycoplasmas is accompanied by prominent lesions in other
organs and/or parts of the body besides the thoracic cavity.
Typical cases of CCPP are characterised by extreme fever (41–43°C), high morbidity and mortality
rates in susceptible herds affecting all ages and both sexes, and abortions in pregnant goats. It
appears to be transmitted by an infective aerosol. After approximately 2–3 days of high fever,
respiratory signs become apparent: respiration is accelerated and painful, and in some cases is
accompanied by a grunt. Coughing is frequent, violent and productive. In the terminal stages,
animals are unable to move – they stand with their front legs wide apart, the neck is stiff and
extended, and sometimes saliva continuously drips from the mouth. Post-mortem examination
reveals fibrinous pleuropneumonia with massive lung hepatisation and pleurisy, accompanied by
accumulation of straw-coloured pleural fluid.
The disease has been shown recently to affect wild ruminants such as the wild goats (Capra
aegagru), Nubian Ibex (Capra ibex Nubian) and Laristan Mouflon (Ovis orientalis laristanica) and
Gerenuk (Litocranius walleri). Clinical disease and seropositivity have been reported in sheep in
contact with affected goats, but the role of sheep as reservoirs of infection is unclear.
Identification of the agent: Definitive diagnosis requires culture of the causative organism from
lung tissue samples and/or pleural fluid taken at post-mortem. After cloning and purification,
isolates can be identified by several biochemical, immunological and molecular tests. Isolating the
causative agent is a difficult task. Recently polymerase chain reaction based tests have been
described and shown to be specific, sensitive and can be applied directly to clinical material, such
as lung and pleural fluid.
Serological tests: Serological tests have been applied for the diagnosis of CCPP in outbreaks in
Eritrea and Turkey. Such tests are best used on a herd basis rather than for diagnosis in individual
animals. The complement fixation test remains the most widely used serological test for CCPP,
although the latex agglutination test is being increasingly used in the diagnostic laboratories as well
as a pen side test; it can used to test whole blood as well as serum. Indirect hemagglutination is
also used. A specific competitive enzyme-linked immunosorbent assay has been developed, but is
not widely available. As with the other serological tests, it does not detect all reactors, but its
specificity and suitability for large-scale testing make it an appropriate test for epidemiological
investigations.
Requirements for vaccines and diagnostic biologicals: Vaccine against CCPP caused by Mccp
is available commercially.
A. INTRODUCTION
Contagious caprine pleuropneumonia (CCPP) is a severe disease of goats occurring in many countries in Africa
and Asia where the total goat population is more than 500 million (Acharya, 1992). The first description of the
disease was in 1873 in Algeria. Shortly after, in 1881, the disease was introduced in the “Cape Colony” of South
Africa by a shipment of Angora goats (Hutcheon, 1881; 1889). The disease was eradicated using a policy of
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Chapter 2.7.6. — Contagious caprine pleuropneumonia
slaughter of the infected goats coupled with a traditional vaccination procedure for the in-contact goats. Classical,
acute CCPP is caused by Mycoplasma capricolum subsp. capripneumoniae (Mccp) (Leach et al., 1993), originally
known as the F38 biotype. This organism was first isolated and shown to cause CCPP in Kenya (MacMartin et al.,
1980; MacOwan & Minette, 1976; 1977a; 1977b); it has subsequently been isolated in the Sudan, Tunisia, Oman,
Turkey, Chad, Uganda, Ethiopia, Niger, Tanzania, Eritrea and the United Arab Emirates. CCPP was first reported
in mainland Europe in 2004, when outbreaks were confirmed in Thrace, Turkey, with losses of up to 25% of kids
and adults in some herds (Ozdemir et al., 2005). However, the exact distribution of the disease is not known and
it may be much more widespread than the zone represented by the countries where Mccp has been isolated as
CCPP is often confused with other respiratory infections and also because the isolation of the causative organism
is difficult.
In CCPP outbreaks in mixed goat and sheep herds, sheep may also be affected, as verified by isolation of Mccp
(Bölske et al., 1995) or detection of antibodies from clinically affected sheep (Kibor & Waiyaki, 1986). Mccp has
also been isolated from healthy sheep (Litamoi et al., 1990) and the role of sheep as a reservoir for the disease
has to be considered.
Recently CCPP was confirmed in wild ruminants kept in a wildlife preservation reserve in Qatar. The disease
affected wild goats (Capra aegagrus), Nubian Ibex (Capra ibex nubiana), Laristan mouflon (Ovis orientalis
laristanica) and Gerenuk (Litocranius walleri) with significant morbidity and mortality in these species (Arif et al.,
2005). Disease indistinguishable from naturally occurring CCPP has been experimentally reproduced with Mccp
by several groups of workers.
B. DIAGNOSTIC TECHNIQUES
The diagnosis of outbreaks of respiratory disease in goats, and of CCPP in particular, is complicated, especially
where it is endemic. It must be differentiated from other similar clinico-pathological syndromes such as: peste des
petits ruminants, to which sheep are also susceptible; pasteurellosis, which can be differentiated on the basis of
distribution of gross lung lesions; and what has been called ‘mastitis, arthritis, keratitis, pneumonia and
septicaemia syndrome or more often as contagious agalactia syndrome (Thiaucourt & Bolske, 1996). As the
longer name implies, the pneumonia is accompanied by prominent lesions in other organs. The disease caused
by Mccp is readily contagious and fatal to susceptible goats of all ages and both sexes, rarely affects sheep and
does not affect cattle.
1.
Identification of the agent
a)
Microscopy of lung exudates, impression smears or sections
Mccp is characterised histopathologically by an interstitial pneumonia with interstitial, intralobular oedema of
the lung (Kaliner & MacOwan, 1976; Nicholas, 2002). It shows a branching filamentous morphology in vivo
that can be observed by dark-field microscopy in exudates or tissue suspensions from lesions or pleural
fluid. Alternatively, smears made from cut lung lesions can be stained by the method of May–Grünwald–
Giesma and examined by light microscopy. The other caprine mycoplasmas show a short filamentous or
coccobacillary morphology. Neither of these techniques provides a definitive diagnosis.
b)
Nucleic acid recognition methods
Two polymerase chain reaction (PCR) assays for the specific identification of Mccp have now been
published. The first one (Bascunana et al., 1994) is based on the amplification of the 16S rRNA gene of the
mycoides cluster. The PCR product is then analysed by restriction enzyme cleavage for the identification of
the Mccp amplicon. The second one (Woubit et al., 2004) is based on a specific amplification. These PCR
techniques can be used directly on clinical materials such as lung tissue and pleural fluid (Bölske et al.,
1996). Due to the difficulty in isolating Mccp, PCR is the technique of choice for the diagnosis of CCPP.
However, isolation of Mccp remains the confirmatory test. All mycoplasmas of the mycoides cluster can be
assigned a precise phylogenetic position by using a multilocus sequence typing approach which may be
used for identification purposes (Manso-Silván et al., 2007).
c)
Gel precipitin tests to detect antigen in tissue specimens
Mccp releases an antigenic polysaccharide (Rurangirwa et al., 1987c) to which a specific monoclonal
antibody (MAb) (WM-25) has been produced (Rurangirwa et al., 1987d; 1992). This MAb
immunoprecipitates in agar gel with the polysaccharide produced by Mccp, and is used to identify the
causative agent in cases of CCPP, particularly when specimens are no longer suitable for culture because of
deterioration during transit.
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Chapter 2.7.6. — Contagious caprine pleuropneumonia
d)
Isolation of mycoplasmas
i)
Selection of samples
The necropsy samples of choice are lung lesions, particularly from the interface between consolidated
and unconsolidated areas, pleural fluid, and mediastinal lymph nodes. If microbiological examination
cannot be performed immediately, samples or whole lungs can be stored at –20°C for considerable
periods (months) with little apparent loss of mycoplasma viability. During transport, samples should
always be kept as cool as possible, as mycoplasma viability diminishes rapidly with increasing
temperature. Lung samples can be dispatched to other laboratories in frozen form.
ii)
Treatment of samples
Swabs are suspended in 2–3 ml of culture medium. Tissue samples are best minced using scissors,
and then shaken vigorously, or pulverised in medium1 using 1 g of tissue to 9 ml of medium. Tissues
should not be ground. The suspension is usually prepared with a mycoplasma medium, but if parallel
bacteriological examination is to be carried out, a high quality bacteriological medium, such as nutrient
broth, may be used to provide a suspension suitable for both forms of examination. Pleural fluid, or a
tissue suspension or swab, is serially diluted through at least three tenfold steps (to a nominal 10–4) in
the selected mycoplasma medium. Dilutions should also be plated on to solid medium.
iii)
Mycoplasma media
The medium used by MacOwan & Minette to culture Mccp organisms (MacOwan & Minette, 1976), is
termed ‘viande foie goat’ (VFG), and includes goat-meat liver broth and goat serum. Alternative
suitable media are WJ (Jones & Wood, 1988), modified Hayflick’s, and modified Newing’s tryptose
broth (Kibor & Waiyaki, 1986) Media enriched with 0.2% (or up to 0.8%) sodium pyruvate perform
considerably better, both for primary isolation and antigen production of Mccp (Miles & Wadher, 1990;
Mohan et al., 1990). Examples of suitable media are as follows (Bölske et al., 1996; Thiaucourt &
Bolske, 1996; Thiaucourt et al., 1992):
 CCPP medium
A. Autoclaved portion (121°C for 15 minutes): Bacto PPLO (pleuropneumonia-like organisms) broth
without crystal violet (Difco) (21 g); deionised water (700 ml).
B. Membrane-filtered portion: Horse (alternatively pig or donkey) serum inactivated at 56°C for
30 minutes (200 ml); fresh yeast extract (100 ml); glucose (sterile solution 0.5 g/ml) (2 ml); and sodium
pyruvate (sterile solution 0.5 g/ml) (8 ml).
Part B is added to A aseptically. Ampicillin (0.1 g/litre) and thallium acetate (250 mg/litre) can be added
to prevent contamination in primary isolations. The final pH of the medium should be 7.4–7.6.
 Modified CCPP medium
A. Autoclaved portion (121°C for 15 minutes): Bacto PPLO broth without crystal violet (Difco) (17.5 g);
glass distilled water (650 ml).
B. Membrane-filtered portion: Horse (alternatively pig or donkey) serum inactivated at 56°C for
30 minutes (250 ml); fresh yeast extract (100 ml); 50% glucose (4 ml); 25% sodium pyruvate (8 ml); 5%
thallium acetate (4 ml); ampicillin (250 mg); and 0.5% phenol red (4 ml). The pH is adjusted to 7.8 with
sodium hydroxide or hydrochloric acid. Part B is added to A aseptically.
Modified Newing’s tryptose broth (Kibor & Waiyaki, 1986) and agar plates (Gourlay’s medium)
(Gourlay, 1964) are routinely used for isolation and maintenance of Mccp in Kenya.
iv)
Medium production, storage and quality control
Certain medium components, particularly serum, yeast extract and deionised water, should be
regularly monitored for growth-promoting capacity before incorporation into mycoplasma media. Lowpassage field isolates should be used for this screening purpose.
Broth media may be stored for at least 6 months at –25°C before use, but penicillin or its analogues
should not be added until final dispensing. Broth media are dispensed into bijoux (1.8 ml or 2.7 ml) or
screw-capped tubes (4.5 ml), and stored for up to 3 weeks at 4°C. Solid media are best made with
agarose (0.9% [w/v]), Noble agar (1.5% [w/v]), or purified agar (0.6% [w/v]). Plates, which are poured
to a depth of 6–8 mm, should be as fresh as possible when used, and should be stored for no more
than 2 weeks at 4°C before use. All culture media should be subjected to quality control and must
1
For example, with the Stomacher 80, A.J. Seward, London, United Kingdom (UK).
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Chapter 2.7.6. — Contagious caprine pleuropneumonia
support the growth of Mycoplasma spp. from small inocula. The reference stain should be cultured in
parallel with the suspicious samples to ensure that the tests are working correctly.
v)
Cultivation
Cultures are incubated at 37°C. Plates are best incubated in a humid atmosphere of 5% CO2, 95% air
or N2, or in a candle jar with a moisture source.
Broth cultures are examined daily for evidence of growth – colour change and the appearance of
floccular material. Gross turbidity indicates bacterial contamination; cultures showing this should be
passed through a 0.45-µm membrane filter before subculture. Broth cultures are subcultured by
inoculation of fresh broth medium with one-tenth of their volume, or by streaking agar medium with a
loop.
Plate cultures are examined every 1–3 days using a stereo microscope (×5–50 magnification) and
transmitted and incident light sources. If negative, the plates are discarded after 14 days. Subculture is
carried out by the transfer of excised agar blocks bearing isolated colonies to either agar (on which the
blocks are pushed, face down) or broth media. Alternatively, an agar plug bearing one colony is drawn
into a Pasteur pipette and discharged into fresh broth medium.
Cloning and purification of isolates is performed by repeated transfer of single colonies representing
each morphological type seen. Colony morphology varies with the medium used, the mycoplasma
species, its passage level and the age of the culture.
In early passage, many mycoplasma species, including M. capricolum subsp. capricolum (Mcc),
produce colonies of bizarre morphology, often small, centreless, and of irregular shape. This effect is
often associated with the use of marginally suitable medium. With passage, such isolates demonstrate
conventional ‘fried egg’ colony morphology, except M. ovipneumoniae, which retains centreless
colonies. Colonies of M. mycoides subsp. mycoides Large Colony (MmmLC) and Mcc may be up to
3 mm in diameter.
Filtration of broth cultures through 0.45 µm filters before subculture aids purification by excluding cell
aggregates.
Cultures suspected of being L-forms of bacteria should be examined for reversion to bacterial form by
three to five passages on solid mycoplasma medium from which antibiotics and thallium acetate have
been omitted.
Broth media used for primary isolation and which have shown no indication of growth by 7 days, should
be subcultured blind.
Cultures of each sample, including one blind subculture, should be examined for a minimum of
3 weeks before being discarded. Titrations in broths, if performed in full (to 10–10), are also read at 3–
4 weeks and are expressed as colour-changing units per transfer volume. Growth on plates is
expressed as colony-forming units (CFU) per ml.
e)
Identification of mycoplasmas
i)
Polymerase chain reaction
Once the organism has been cultured, verification of Mccp can be achieved in 1 day by PCR. There
are now various PCR tests that can be applied for an identification of Mccp strains. The first test
(Bascunana et al., 1994) is based on amplification of a segment of the 16S rRNA gene. The amplified
fragment is common to the mycoides cluster. However, when the amplicon is digested with
endonuclease PstI, a unique cleavage pattern of three fragments for Mccp is observed when the
enzyme digests are analysed in agarose gel electrophoresis and stained with ethidium bromide (Bölske
et al., 1996). The second test is based on a specific amplification and can allow confirmation of CCPP
within a few hours (Woubit et al., 2004).
Recently, PCR and sequencing has been used to establish the molecular epidemiology of CCPP.
These tests can be performed on dried samples, such as pleural fluid on filter papers. The sequencing
allows a precise identification of the species (the cleavage site for the 16S rRNA and a specific
detection for the ‘locus H2’) (Lorenzon et al., 2002; Pettersson et al., 1998).
Identification of Mccp strains by PCR (and sequencing) has now superseded all other techniques
because of its rapidity and reliability. However PCR reactions must be performed with great care to
prevent contaminations.
ii)
Biochemical tests
Wild strains should be passaged, and preferably cloned, three times before identification is attempted.
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Biochemical tests cannot identify an isolate unequivocally, which at present can only be done by
serological or genetic means. Intraspecies variation in some biochemical reactions is often
considerable (Houshaymi et al., 2000), but some tests perform a useful function both as a preliminary
screening system and in providing supportive data for serological findings.
The tests most commonly used are glucose breakdown, arginine hydrolysis, ‘film and spots’ formation,
reduction of tetrazolium chloride (aerobically and anaerobically), phosphatase activity, serum digestion
and digitonin sensitivity. The first three of these tests are performed routinely in isolation and cultivation
procedures. Glucose breakdown is indicated by acid (yellow) changes, and arginine hydrolysis by
alkaline (red) changes in broth media, using phenol red as indicator. Arginine use cannot be assessed
on conventional medium for isolation and culture as the media for testing the arginine deiminase
pathway should contain high concentrations of arginine and no glucose. Film and spots’ describes an
apparent wrinkling of the agar surface due to the deposition on it of an iridescent film of lipid, together
with the development of black spots within the medium in the vicinity of ageing colonies. This
phenomenon, produced by three mycoplasmas of small ruminants, is demonstrated on agar media
containing 20% or more serum, preferably of horse or pig origin. Supplementation of the medium with
10% egg yolk emulsion improves the sensitivity of the test.
The remaining biochemical tests require specific media or reagents. The test for tetrazolium reduction
provides corroborative evidence of the mycoplasmal nature of M. agalactiae isolates, as this organism is
neither glycolytic nor arginine-hydrolysing. Serum digestion (Freundt, 1983) distinguishes members of
ruminant mycoplasmas, and phosphatase production (Bradbury, 1983) separates Mcc from other members
of this cluster. Digitonin sensitivity distinguishes members of the order Mycoplasmatales from those of the
order Acholplasmatales (Freundt et al., 1973). A diagnostic medium is available that enables the specific
detection of Mccp growing on agar medium: colonies show a red coloration (Ozdemir et al., 2005).
iii)
Serological identification
Mycoplasmal antigens used in hyperimmune serum production are often contaminated with medium
constituents. The antibodies stimulated by these contaminants can cause false-positive reactions in
serological identification tests. This problem is avoided by absorption of the antiserum with the medium
used to produce the antigen (10 mg lyophilised medium per ml of antiserum), or by growing the
mycoplasmas to be used as antigens in medium containing homologous animal components, e.g.
growth in VFG medium to immunise goats.
Because of the close serological relationships between members of the ‘M. mycoides cluster’, isolates from
cases of CCPP should, preferably, be identified by at least two of the three tests described below.

Growth inhibition test
The growth inhibition test (GIT) is the simplest and most specific, but the least sensitive, of the tests
available. It depends on the direct inhibition of growth on solid medium by specific hyperimmune serum, and
detects primarily surface antigens (Dighero et al., 1970).
Mccp appears to be highly homogeneous serologically and wide zones of inhibition free of ‘breakthrough’
colonies are observed with antiserum to the type strain, regardless of the source of the test strain (Jones &
Wood, 1988). Mccp cross-reacts with Leach’s bovine group 7 (PG50), M. equigenitalium and M. primatum in
the GIT when polyclonal antisera are used, but an MAb specific for Mccp in the GIT has been produced
(Rurangirwa et al., 1987d). The MAb reagent, WM25, has been reported to be specific for (Mccp) isolates by
the disc growth inhibition method, which will exclude M. agalactiae, Mcc and the other members of the
‘M. mycoides cluster’ associated with goats, but not bovine group 7 (not usually found in goats): the latter
can be excluded, however, by colony indirect fluorescence tests (Belton et al., 1994). A small proportion of
Mccp isolates also cross-react in the GIT with antiserum to Mcc. Group seven of Leach strains can
sometimes be found in goats although it is rare. Results should be interpreted carefully as some bovine
strains have been misidentified by the GIT using the ‘specific’ antiserum.

Test procedure
i)
Broth culture in mid-to-late logarithmic phase is used at three tenfold dilutions, the selection of which is
related to the vigour of growth of the isolate on agar.
ii)
Agar plates are dried for 30 minutes at 37°C.
iii)
Sterile paper disks of 6–7 mm in diameter are impregnated with a drop (10–20 µl) of undiluted
antiserum. Disks may be used wet, in which form they can be stored at –20°C, or they can be
lyophilised (Dighero et al., 1970), which allows storage at 4°C.
iv)
Using a separate plate for each dilution of culture, 1 ml or 2.5 ml volumes are pipetted on to 5 cm or
10 cm diameter plates, respectively. The inoculum is dispersed evenly over the plate, then the excess
is removed.
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v)
The plates are dried at 20–30°C for 15–20 minutes, preferably under a protective hood, until no visible
liquid is present on the surface. Sufficient residual moisture should remain to enable freeze-dried disks
to adhere to the agar surface.
vi)
Several disks, each impregnated with a different antiserum (selected on the basis of sample source
and the biochemical reactions and colony morphology of the isolate), are carefully placed on the agar
plates; isolates from CCPP cases should be screened with antisera against Mccp, MmmLC, Mcc,
M. mycoides subsp. capri (Mcc) and M. ovipneumoniae. A disk containing 1.5% digitonin should also
be included on the plates.
vii)
The plates are incubated at 37°C for 2–6 days. Initial overnight incubation at 27°C can increase the
sensitivity of the test. Inhibition by digitonin is generally readily apparent; however, inhibition by
antiserum may be more difficult to interpret, with suppression rather than total inhibition of growth,
depending on the species of mycoplasma, colony density and potency of the antiserum. ‘Breakthrough’
colonies are commonly observed within zones of inhibition. Circular precipitin bands are occasionally
seen around disks. Positive inhibition is regarded as a zone of 2 mm or more.

Growth precipitation test
The growth precipitation test detects soluble cytoplasmic and extramembranous antigens released by
growing cultures and allowed to diffuse through solid mycoplasma growth medium towards mycoplasma
antiserum during growth (Krogsgaard-Jensen, 1972). As with the gel precipitin test, there are strong crossreactions within the mycoides cluster. If growth inhibition is performed using MAb WM252, which is specific
for Mccp, both specific inhibition and a growth precipitin line are achieved simultaneously.

Indirect fluorescence antibody test
The direct and indirect fluorescent antibody tests are the most effective of the various serological methods
for identifying most mycoplasmas (Rosendal & Black, 1972). They are simple, rapid, and sensitive, yet
economical in the use of antiserum. Several forms have been described, the most commonly used and
perhaps best being the indirect fluorescent antibody (IFA) test applied to unfixed colonies on agar.
Antiserum against a single strain is sufficient to identify field isolates of that species, and antisera are diluted
before use. Cultures do not have to be cloned, but the test is usually applied only after several passages
have indicated whether the culture contains more than one species and the growth characteristics of the
organism(s) present.

Test procedure
i)
Two agar plates are predried at 37°C for 30 minutes. Each one is flooded with a different dilution of test
broth culture, the dilutions being selected according to the vigour of growth of the strain on agar
medium. Alternatively, a drop of undiluted culture is spread over a single 5 cm plate using an L-shaped
glass rod.
ii)
The plates are incubated at 37°C until the first evidence of growth is observed. If the IFA test cannot be
performed immediately, the plates can be stored at 4°C for up to 4 weeks.
iii)
Several blocks of approximately 0.5–1 cm2 are excised from areas where colonies are numerous, but
not confluent. The blocks of each agar culture are cut to the same geometric shape to enable
recognition of origin, a different shape being used for each isolate. Several blocks of each isolate are
distributed (colony surface facing upwards) on to several different slides, each slide being used for a
different mycoplasma antiserum. The colony surface of each block is identified for future reference by
undercutting one corner.
iv)
Rabbit anti-mycoplasma (ra-m) serum or normal rabbit serum (NRS; as a control on a duplicate block)
at a suitable dilution in normal saline or phosphate buffered saline (PBS), pH 7.2, is gently pipetted on
to each agar block until the surface area is totally covered. The optimal dilution of ra-m is determined
by chequerboard titration against the fluorescein isothiocyanate (FITC)-conjugated anti-rabbit
immunoglobulin serum (a-r lg-FITC) used.
v)
The flooded blocks are incubated on their slides at room temperature for 30 minutes in a humid
chamber.
vi)
All blocks on one slide are tipped into a 10 ml tube containing approximately 7 ml of PBS.
vii)
The plugged tubes are rotated at 18–30 rpm for 10 minutes. The PBS is then decanted and replaced
with fresh PBS, and the tubes are rotated again for 10 minutes.
viii) The PBS is decanted and the blocks are placed colony surface facing upwards on their respective
slides. Excess moisture is blotted off.
2
Available from Kenya Agricultural Research Institute (KARI), P.O. Box 57811, Nairobi, Kenya.
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ix)
All blocks are flooded with a-r lg-FITC at optimal dilution.
x)
The blocks are incubated again for 30 minutes at room temperature in a humid chamber, then tipped
into tubes containing fresh PBS, and washed twice by rotating, as before.
xi)
The blocks, replaced colony surface facing upwards on their respective slides, are examined by an epiimmunofluorescent microscope using the settings recommended by the manufacturer for FITC.

Notes on the indirect fluorescent antibody test
xii)
Working dilutions of ra-m and a-r lg-FITC should be kept at 4°C, which limits their shelf life to
approximately 1 week.
xiii) Isolates from CCPP should be examined using antisera against Mccp, MmmLC, Mmc (M. mycoides
subsp. capri) and Mcc, positive control cultures should comprise their type strains, namely Mccp, Y
goat, PG3 and California kid, respectively.
xiv) A negative (NRS-treated) control should always be incorporated for each culture.
xv)
f)
Interpretation of the IFA test can be difficult. Autofluorescence is produced by some species,
particularly acholeplasmas. Even in pure cultures, a proportion of colonies may not stain positively with
the relevant antiserum; this is particularly true of Mcc. Otherwise, poor results are usually ascribable to
the use of an agar culture that has been allowed to grow for too long, or to the use of antiserum that
has deteriorated with dilution and age.
Other identification tests
Metabolism inhibition (Taylor-Robinson et al., 1966) and tetrazolium reduction inhibition (Senterfit & Jensen,
1996) are other tests sometimes used in the identification of caprine mycoplasmas. A gene probe, F38-12,
capable of distinguishing Mccp has been developed (Taylor et al., 1992).
A polysaccharide-specific antigen detection latex agglutination test has been developed to detect CCPP
antigen (March et al., 2000). In this test, latex beads are coated with polyclonal immunoglobulin IgG (rabbit)
directed against Mccp polysaccharide and used to detect the antigen in the serum of goats with CCPP. This
test is proposed to be inexpensive, easy to carry out in the field and useful for detecting CCPP in its earliest
stages.
2.
Serological tests
Serology has not been widely applied to identifying the cause of outbreaks of pleuropneumonia in goats and
sheep. Endemic infections with MmmLC and Mmc can produce a background of positive titres to these organisms
in a proportion of apparently healthy animals (Jones & Wood, 1988), and under experimental conditions
seroconversion to M. mycoides can occur in goats with no clinical signs of disease. Acute cases caused by Mccp
rarely show positive titres to the organism before death (MacOwan & Minette, 1977b; Muthomi & Rurangirwa,
1983; Thiaucourt & Bolske, 1996), perhaps because antibodies are ‘eclipsed’ by circulating mycoplasma antigens
(Muthomi & Rurangirwa, 1983). Seroconversion to Mccp in experimentally infected animals is observed, by the
complement fixation (CF) test and indirect haemagglutination (IHA) test, to start 7–9 days after the appearance of
clinical signs, to peak between days 22 and 30, and to decline rapidly thereafter (Muthomi & Rurangirwa, 1983).
These various observations indicate that serology should be applied on a herd, not an individual basis, and that
whenever possible, paired serum samples collected 3–8 weeks apart, should be examined.
a)
Complement fixation test (the prescribed test for international trade) (MacOwan & Minette, 1976)
The CF test in various forms remains the most widely used serological test for diagnosis of contagious
bovine pleuropneumonia (Gourlay, 1983; Perreau et al., 1976). In CCPP, the CF test was used for detection
of Mccp infection (MacOwan & Minette, 1976) and it has been found to be more specific, though less
sensitive, than the IHA test (Muthomi & Rurangirwa, 1983). Its main disadvantage is the high level of
technical expertise required to perform the test (Gourlay, 1983).
One method of performing the test is as follows. To prepare the antigen, 2 litres of culture of titre higher than
109 CFU/ml is centrifuged at 40,000 g for 1 hour at 5°C. The deposit is resuspended and washed three
times in physiological saline prior to storage in 0.5–1.0 ml volumes at –20°C.
Sterile broth treated as above constitutes sediment antigen, and a freeze-dried broth reconstituted at
200 mg/ml constitutes a second control antigen. Prior to testing, the antigen is diluted 1/60 and
ultrasonicated for 3 minutes at low power in a container of iced water. The sonicate is centrifuged at 1250 g
for 30 minutes to remove any debris, and stored at –20°C. If stored for more than 2–3 weeks the antigen
should be recentrifuged.
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
Test procedure
Microtitre plate tests are performed using 0.025 ml volumes, two volumes containing three mean haemolytic
doses of complement, and a 1.5% (v/v) final concentration of sheep red blood cells (SRBCs) in U-bottomed
microtitre plates as follows:
i)
The following are mixed and incubated at 37°C for 45 minutes:

25 µl of doubling dilutions of test serum (heat inactivated at 56°C for 30 minutes) starting with
½ dilution;

25 µl of antigen (the dilution of the antigen must be determined in a chequerboard titration using a
known positive serum);

25 µl of complement (3 haemolytic units).
ii)
25 µl of sensitised SRBCs, at a final concentration of 1.5% (v/v), is mixed and the plates are incubated
at 37°C for 45 minutes.
iii)
The plates are incubated at 4°C for 1 hour to allow the unlysed cells to settle.
iv)
Reading the results: The titre will be the highest serum dilution that will fix 50% of the complement, i.e.
50% haemolysis.
•
Controls
In all CF tests a number of controls are required:
b)
i)
Indicator systems (RBCs + haemolysin) alone to ensure that RBCs do not lyse spontaneously.
ii)
Indicator system with complement only to show that enough complement is present to lyse the cells.
iii)
Indicator system with antigen only and no complement to show that antigen alone does not lyse the
cells.
iv)
Indicator system with serum alone and no complement to show that the serum alone does not lyse the
cells.
v)
Indicator system with complement and antigen to detect any anticomplementary activity of the antigen.
vi)
Indicator system with the complement and serum to detect any anticomplementary activity of the
serum.
Latex agglutination test
Latex beads sensitised with the polysaccharide produced by Mccp and present in culture supernatant have
been used in a slide agglutination test (Houshaymi et al., 2002; Rurangirwa et al., 1987a). This test is
presently used routinely in Kenya. It is a very useful test in an outbreak because it can be performed at the
penside using a drop of whole blood.
Both CF test and IHA test findings emphasise the difficulties inherent in the serological diagnosis of CCPP
when using whole cell or membrane preparations as antigen. The use of the more defined antigen, the
polysaccharide elaborated by Mccp, provides greater specificity, as there is no cross-reactivity with sera
against the other three principal caprine mycoplasmas.
c)
Competitive enzyme-linked immunosorbent assay
A competition ELISA has been developed (Thiaucourt et al., 1994) and proved both specific and sensitive.
However due to erratic and low demand the production of this test has been stopped until a new format is
developed to ensure longer stability of reagents.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
The first experimental vaccine against Mccp was a live high passage Mccp (MacOwan & Minette, 1978). When
inoculated intratracheally, it proved innocuous and protected goats against experimental challenge. However,
more recent work has concentrated on inactivated forms of vaccine. The current form used in Kenya (where
inactivated Mccp, vaccines have been in use for several years) contains inactivated Mccp suspended in saponin,
has been described as having a shelf life of at least 14 months and the optimal dose of 0.15 mg provides
protection for over 1 year (Rurangirwa et al., 1987b).
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1.
Seed management
a)
Characteristics of the seed
The master seed was isolated from the lungs of a sick goat. In Kenya the strain used is called ‘Yatta’ and it
has been confirmed to be Mccp by GIT and PCR, after 15 passages in culture. The master seed should also
be pure and free from other contaminants.
b)
Method of culture
The master seed is established and stored in a freeze-dried form in 1 ml ampoules. The working seed is
prepared by amplifying the master seed in modified Newing’s broth to make a bulk of 4 litres. Its growth is
arrested at the growth phase before filament formation.
This culture is tested for sterility before it is distributed in aliquots of 20 ml volumes and stored frozen at
–20°C.
c)
Validation as a vaccine
The master seed should be prepared from lung specimens or pleural fluids of a goat that dies of pneumonia,
showing all the clinical signs of CCPP. The isolate must be confirmed by GIT or by PCR to be Mccp. It must
be checked for sterility, safety, potency and extraneous agents.
2.
Method of manufacture
For vaccine production, a working seed is first established by amplifying an aliquot of freeze-dried master seed in
modified Newing’s broth (Kibor & Waiyaki, 1986), to make a bulk of 4 litres of culture. This culture is aliquoted in
20 ml volumes and frozen at –20°C. The vaccine is grown in 5 litre pots containing 4 litres of modified Newing’s
broth. Each pot is sampled aseptically for sterility testing before 20 ml of working seed is inoculated into each
4 litres of medium. These pots are incubated at 37°C for 4–6 days depending on how fast the mycoplasma grows.
After the filaments sediment, the antigen is harvested by centrifugation at 2600 g from pots that are not
contaminated. (The filaments are thin and white and sometimes unite to form a resemblance of an inverted pine
tree. This occurs on days 4–6 and it is at this point that they become heavy and sediment.) The pellet is
resuspended in sterile normal saline and centrifuged to remove the remnants of the growth medium. The pellet is
resuspended again in a small volume of sterile normal saline to make a viscous suspension.
Saponin is added to inactivate the mycoplasma at 3 mg saponin to 1 ml of antigen. It is left agitating overnight at
4°C using a magnetic stirrer.
NOTE: Saponin also acts as an adjuvant. Three samples are taken aseptically from this suspension and tested
for sterility, for protein estimation and for the innocuity test.
3.
In-process control
During production, the following tests are carried out to ensure that the product remains pure and safe. These
tests are carried out by quality assurance staff. Each pot is sampled aseptically before inoculation to test for
sterility.
After maturity of culture, only pots that have not shown signs of contamination are pooled for centrifugation; the
contaminated pots are decontaminated and discarded after autoclaving.
After inactivation with saponin, a sample is taken aseptically for sterility, another one for innocuity and another
one for the protein estimation test.
a)
Sterility test
This test is aimed at verifying the absence of fungal and bacterial contaminants. Two tubes of thioglycollate
are inoculated with 1 ml of the sample each. The tubes contain about 15 ml of broth. These are incubated at
37°C to eliminate aerobics, microaerophilics and anaerobics. The other medium used is soybean casein
digest broth. Four tubes are inoculated with 12 ml of sample each.
Two tubes are incubated at 37°C and another two at room temperature (25°C) to eliminate bacterial and
fungal contaminants. All media are incubated for 14 days with controls. Absence of any growth shows that
the sample is not contaminated.
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b)
Innocuity test
This test aims to demonstrate the absence of living mycoplasma in the vaccine: 0.3 ml of inactivated antigen
is added to 2.7 ml of modified Newing’s broth in a tube and tenfold dilutions are made from 10–1 to 10–9. A
positive control is set up using a viable culture of M. capripneumoniae and a negative control using three
uninoculated tubes of medium. All are incubated at 37°C for 12 days. If there is no growth in the tubes
inoculated with test sample, the vaccine passes the test.
c)
Protein content estimation
The protein content is measured by the bicinchoninic acid method by comparison with a bovine albumin
standard.
4.
Batch control
a)
Sterility
The sterility test on the final batch is carried out as described in Section C.3.a except that four bottles of
each batch are pooled together and samples of the pool are used.
b)
Safety
Every batch of CCPP vaccine must be proven to be safe in laboratory animals. Two guinea-pigs are each
inoculated by the intramuscular route in the hind leg and another two guinea-pigs are inoculated by the
peritoneal route with 0.5 ml containing five doses of vaccine.
If the vaccine is safe, the guinea-pigs should not show any sign of disease for a 14-day observation period
and, on post-mortem, there should be no abscesses on the site of inoculation and in the peritoneal cavity,
respectively. If any vaccine-related deaths occur during the observation period, the vaccine fails. If on postmortem examination, abscesses are observed at the site of inoculation and the peritoneum, the vaccine also
fails.
c)
Potency
Research is in progress at the Kenya Agricultural Research Institute, Muguga, Kenya, to develop a test for
potency.
d)
Duration of immunity
The vaccine protects goats for 14 months. It is, however, recommended to boost immunity after 1 year.
e)
Stability
If stored at 4°C, the vaccine has a shell life of 1 year. Before use the vaccine should be shaken thoroughly,
for even distribution of antigen.
f)
Preservatives
At present preservatives are not used in the vaccine.
g)
Precautions
Side-effects of the CCPP vaccine include development of a swelling at the site of inoculation and fever for
1–2 days following vaccination, accompanied with inappetance. The swelling may last between 1 and
14 days and is due to the saponin adjuvant. Accidental self-injection causes severe irritation.
5.
Tests on the final product
a)
Safety
Every batch of vaccine should be tested for safety in laboratory animals as described in C.4.b.
b)
Potency
Once the potency test has been developed, every batch of vaccine will be required to be tested for potency.
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KALINER G. & MACOWAN K.J. (1976). The pathology of experimental and natural contagious caprine
pleuropneumonia in Kenya. Vet. Med. [B], 2, 652–661.
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KROGSGAARD-JENSEN A. (1972). Mycoplasma: growth precipitation as a serodiagnostic method. Appl. Microbiol.,
23, 553–558.
LITAMOI J.K., WANYANGU S.W. & SIMAM P.K. (1990). Isolation of Mycoplasma biotype F38 from sheep in Kenya.
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LEACH R.H., ERNO H. & MACOWAN K.J. (1993). Proposal for designation of F38-type caprine mycoplasmas as
Mycoplasma capricolum subsp. capripneumoniae subsp. nov. and consequent obligatory relegation of strains
currently classified as M. capricolum (Tully, Garile, Edward, Theodore & Erno, 1974) to an additional new
subspecies, M. capricolum subsp. capricolum subsp. nov. Int. J. Syst. Bacteriol., 43, 603–605.
MACMARTIN D.A., MACOWAN K.J. & SWIFT L.L. (1980). A century of classical contagious caprine pleuropneumonia:
from original description to aetiology. Br. Vet. J., 136, 507–515.
MACOWAN K.J. & MINETTE J.E. (1976). A mycoplasma from acute contagious caprine pleuropneumonia in Kenya.
Trop. Anim. Health Prod., 8, 91–95.
MACOWAN K.J. & MINETTE J.E. (1977a). The role of mycoplasma strain F38 in contagious caprine
pleuropneumonia in Kenya. Vet. Rec., 101, 380–381.
MACOWAN K.J. & MINETTE J.E. (1977b). Contact transmission of
pleuropneumonia (CCPP). Trop. Anim. Health Prod., 9, 185–188.
experimental contagious
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MACOWAN K.J. & MINETTE J.E. (1978). The effect of high passage mycoplasma strain F38 on the courses of
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Mycoplasma capricolum subsp. capripneumoniae capsular polysaccharide-specific antigen detection latex
agglutination test. J. Clin. Microbiol., 38, 4152–4159.
MANSO-SILVÁN L. PERRIER X. & THIAUCOURT F. (2007). Phylogeny of the Mycoplasma mycoides cluster based on
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from the lungs of Nigerian goats. Zentralbl. Bakteriol. (Suppl.), 20, 841–843.
MUTHOMI E.K. & RURANGIRWA F.R. (1983). Passive haemagglutination and complement fixation as diagnostic tests
for contagious caprine pleuropneumonia caused by the F38 strain mycoplasma. Res. Vet. Sci., 35, 1–4.
NICHOLAS R.A.J. (2002). Contagious caprine pleuropneumonia. Recent Advances in Goat Diseases, Tempesta
M., ed., International Veterinary Information Service, Ithaca, NY, USA.
www.ivis.org/advances/Disease_Tempesta/nicholas/chapter_frm.asp.
OZDEMIR U., OZDEMIR S., MARCH J., CHURCHWOOD C. & NICHOLAS R.A.J. (2005). Outbreaks of CCPP in the Thrace
region of Turkey. Vet. Rec., 156, 286–287.
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ruminants: un test de fixation du complement. Bull. Acad. Vet. Fr., 49, 185–192.
PETTERSSON B., BOLSKE G., THIAUCOURT F., UHLEN M. & JOHANSSON K.K. (1998). Molecular evolution of
Mycoplasma capricolum subsp. capripneumoniae strains based on polymorphisms in the 16S rRNA genes. J.
Bacteriol., 180, 2350–2358.
ROSENDAL S. & BLACK F.T. (1972). Direct and indirect immunofluorescence of unfixed and fixed mycoplasma
colonies. Acta Pathol. Microbiol. Scand., 80, 615–622.
RURANGIRWA F.R., MCGUIRE T.C., KIBOR A. & CHEMA S. (1987a). A latex agglutination test for field diagnosis of
caprine pleuropneumonia. Vet. Rec., 121, 191–193.
RURANGIRWA F.R., MCGUIRE T.C., KIBOR A. & CHEMA S. (1987b). An inactive vaccine for contagious caprine
pleuropneumonia. Vet. Rec., 121, 397–402.
RURANGIRWA F.R., MCGUIRE T.C., MAGNUSON N.S., KIBOR A. & CHEMA. S. (1987c). Composition of polysaccharide
from mycoplasma (F-38) recognized by antibodies from goats with contagious caprine pleuropneumonia. Res.
Vet. Sci., 42, 175–178.
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RURANGIRWA F.R., MCGUIRE T.C., MUSOKE A.J. & KIBOR A. (1987d). Differentiation of F38 mycoplasmas causing
contagious caprine pleuropneumonia with a growth-inhibiting monoclonal antibody. Infect. Immun., 55, 3219–
3220.
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International Organization for Mycoplasmology, August 2–7 1992, Ames, Iowa, USA. IOM Letters, Vol. 2 FP5/2, p.
128.
SENTERFIT L.B. & JENSEN K.E. (1996). Antimetabolic antibodies to Mycoplasma pneumoniae measured by
tetrazolium reduction inhibition. Proc. Soc. Exp. Biol. Med., 122, 786–790.
TAYLOR T.K., BASHRUDDIN J.B. & GOULD A.R. (1992). Relationship between members of the Mycoplasma mycoides
cluster as shown by DNA probes and sequence analysis. Int. J. Syst. Bacteriol., 42, 593–601.
TAYLOR-ROBINSON D., PRUCELL R.H., WONG D.C. & CHANOCK R.M. (1966). A colour test for the measurement of
antibody to certain mycoplasma species based on the inhibition of acid production. J. Hyg. (Camb.), 64, 91–104.
THIAUCOURT F. & BOLSKE G. (1996). Contagious caprine pleuropneumonia and other pulmonary mycoplasmoses of
sheep and goats. Rev. sci. tech. Off. Int. Epiz., 15, 1397–1414.
THIAUCOURT F., BOLSKE G., LIBEAU G., LE GOFF C. & LEFEVRE P.-C. (1994). The use of monoclonal antibodies in the
diagnosis of contagious caprine pleuropneumonia (CCPP). Vet. Microbiol., 41, 191–203.
THIAUCOURT F., GUERIN C., MADY V. & LEFEVRE P.-C. (1992). Diagnostic de la pleuropneumonie contagieuse
caprine: améliorations récentes. Rev. sci. tech. Off. Int. Epiz., 11, 859–865.
WOUBIT S., LORENZON S., PEYRAUD A., MANSO-SILVAN L. & THIAUCOURT F. (2004). A specific PCR for the
identification of Mycoplasma capricolum subsp. capripneumoniae, the causative agent of contagious caprine
pleuropneumonia (CCPP). Vet. Microbiol., 104, 125–132.
*
* *
NB: There is an OIE Reference Laboratory for Contagious caprine pleuropneumonia
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for contagious caprine pleuropneumonia
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CHAPTER 2.7.7.
ENZOOTIC ABORTION OF EWES
(ovine chlamydiosis)
SUMMARY
Ovine chlamydiosis, also known as enzootic abortion of ewes (EAE) or ovine enzootic abortion
(OEA), is caused by the bacterium Chlamydophila abortus. Chlamydial abortion typically occurs in
the last 2–3 weeks of pregnancy with the appearance of stillborn lambs and inflamed placentas.
However, infection can also result in the delivery of full-term stillborn lambs or weak lambs that do
not survive longer than 48 hours. Infected ewes can also give birth to healthy lambs and it is not
uncommon to observe delivery of a dead and a weak or healthy lamb. There are rarely any
predictive signs that abortion is going to occur, although behavioural changes and a vulval
discharge can be observed in the last 48 hours of pregnancy.
Diagnosis of enzootic abortion depends on the isolation and identification of the causative agent or
detection of the agent or its nucleic acid in the products of abortion or vaginal excretions of freshly
aborted females. A humoral antibody response may be detected following abortion. Goats as well
as sheep and, less commonly, cattle, pigs, horses and deer, can be affected. Chlamydiosis of small
ruminants is a zoonosis and the organism must be handled with biosafety precautions. Pregnant
women are particularly at risk.
Identification of the agent: The basis for a positive diagnosis of infection with C. abortus depends
on a history of abortion in sheep or goats (often in late pregnancy), evidence of necrotic placentitis,
and the demonstration of large numbers of the organism in stained smears of affected placentae.
The still moist fleece of fetuses or vaginal swabs of females that have freshly aborted are also
useful. Care is needed to distinguish cotyledonary damage caused by Toxoplasma gondii and, in
stained smears, to be aware of the morphological similarities between C. abortus and Coxiella
burnetii, the agent of Q fever.
Chlamydial antigen can be detected by enzyme-linked immunosorbent assay, immunohistochemistry or the fluorescent antibody test, whereas chlamydial DNA can be detected by the
polymerase chain reaction or by microarray. Some of these methods are available in commercial kit
form.
Chlamydophila abortus can be isolated only in living cells; thus facilities for growth in chicken
embryos or cell cultures, with appropriate biohazard containment, are required.
Serological tests: A rise in antibody titre to C. abortus, detected by the complement fixation (CF)
test, is common after abortion or stillbirth, but this does not occur in every case. Chlamydophila
abortus shares common antigens with C. pecorum and some Gram-negative bacteria, so that the
CF test is not wholly specific, nor does it distinguish between responses to vaccination and to
infection. Low CF titres need to be interpreted with caution, particularly if these are encountered in
individual animals or in flocks with no history of abortion.
Alternative serological tests have been developed and some commercialised, but none has been
sufficiently appraised so far for field use. A delayed hypersensitivity reaction to chlamydial antigen
can be elicited in infected sheep, but the procedure is not amenable to routine use.
Requirements for vaccines: Inactivated and live vaccines are available that have been reported to
prevent abortion and to reduce excretion. They assist in control of the disease but will not eradicate
it. Serological screening during the period after parturition helps to identify infected flocks, to which
control measures can then be applied.
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A. INTRODUCTION
Ovine chlamydiosis (enzootic abortion of ewes [EAE] or ovine enzootic abortion [OEA]) is caused by the
bacterium Chlamydophila abortus. Chlamydial abortion in late pregnancy causes serious reproductive wastage in
many sheep-rearing areas of the world, particularly where flocks are closely congregated during the parturient
period (Aitken & Longbottom, 2007; Longbottom & Coulter, 2003). Abortion typically occurs in the last 2–3 weeks
of pregnancy with the appearance of stillborn lambs and grossly inflamed placentas. Infection can also result in
the delivery of full-term stillborn lambs and weak lambs that generally fail to survive beyond 48 hours. It is also not
uncommon in multiple births for an infected ewe to produce one dead lamb and one or more weak or healthy
lambs. Infection is generally established in a ‘clean’ flock through the introduction of infected replacements and
results in a small number of abortions in the first year, which is followed by an ‘abortion storm’ in the second year
that can affect up to around 30% of ewes.
Infected animals show no clinical illness prior to abortion, although behavioural changes and a vulval discharge
may be observed in ewes within the last 48 hours of pregnancy. Pathogenesis commences around day 90 of
gestation coincident with a phase of rapid fetal growth when chlamydial invasion of placentomes produces a
progressively diffuse inflammatory response, thrombotic vasculitis and tissue necrosis. Milder changes occur in
the fetal liver and lung and, in cases in which placental damage is severe, there may be evidence of hypoxic brain
damage (Buxton et al., 2002). Abortion probably results from a combination of impairment of materno-fetal
nutrient and gaseous exchange, disruption of hormonal regulation of pregnancy and induced cytokine aggression
(Entrican, 2002).
Chlamydial abortion also occurs in goats and, less frequently, cattle, pigs, horses and deer may be affected. In
sheep, abortion in late pregnancy with expulsion of necrotic fetal membranes are key diagnostic indicators, with
care being needed to distinguish the diffuse pattern of necrosis from that caused by Toxoplasma gondii
(cotyledons only). Distinction from other infectious causes of abortion such as brucellosis (see chapter 2.7.2),
coxiellosis (see chapter 2.1.12) or other bacterial pathogens (Campylobacter [see chapter 2.9.3], Listeria [see
chapter 2.9.7], Salmonella [see chapter 2.9.9]) can be achieved by microscopy and/or culture.
Taxonomically, the family Chlamydiaceae has been divided into two genera and nine species based on sequence
analysis of the 16s and 23s rRNA genes (Everett et al., 1999). The genus Chlamydia includes C. trachomatis
(humans), C. suis (swine) and C. muridarum (mouse and hamster). The genus Chlamydophila includes C. psittaci
(avian), C.felis (cat), C. abortus (sheep, goat and cattle), C. caviae (guinea-pig), the former species C. pecorum
(sheep and cattle) and C. pneumoniae (humans). A proposal has been mooted to recombine all the species in a
single genus (Chlamydia) but has not been adopted for the purposes of this chapter. The terms ‘chlamydiosis’
and ‘chlamydia(e)’ are used to refer to members of the genus Chlamydia in general. However, a binomial of the
generic and specific names is used when referring to a particular chlamydial species. Infected females shed vast
numbers of infective C. abortus at the time of abortion or parturition, particularly in the placenta and uterine
discharges and at subsequent lambing (Papp et al., 1994), thus providing an infection source in the flock. Aborted
ewes do not usually abort again from C. abortus infection. Recent evidence suggests that the proportion of
infected ewes is reduced at the subsequent breeding season and only low levels of chlamydial DNA are detected
during the periovulation period and at lambing, so that this would not have significant impact on the epidemiology
(Livingstone et al., 2009; Gutierrez et al., 2011).

Zoonotic risk and biosafety requirements
Human infection may be acquired from infected products of abortion or parturition or from carelessly handled
laboratory cultures of the organism, with effects that range from subclinical infection to acute influenza-like illness.
Appropriate precautions should be taken and containment level 2 practices are recommended when handling
cultures and potentially infected tissues (see Chapter 1.1.3 Biosafety and biosecurity in the veterinary
microbiology laboratory and animal facilities). Authenticated cases of human placentitis and abortion caused by
C. abortus of ovine origin indicate that pregnant women are at special risk and should not be exposed to sources
of infection (Longbottom & Coulter, 2003; Sillis & Longbottom, 2011).
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
a)
Smears
Where the clinical history of the flock and the character of lesions in aborted placentae suggest enzootic
abortion, a diagnosis can be attempted by microscopic examination of smears made from affected chorionic
villi or adjacent chorion. Several staining procedures are satisfactory, for example, modified Machiavello,
Giemsa, Brucella differential, or modified Ziehl–Neelsen stains (Stamp et al., 1950). In positive cases
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Chapter 2.7.7. — Enzootic abortion of ewes (ovine chlamydiosis)
stained by the latter method and examined under a high-power microscope, large numbers of small
(300 nm) coccoid elementary bodies are seen singly or in clumps stained red against the blue background of
cellular debris. Under dark-ground illumination, the elementary bodies are pale green. If placental material is
not available, smears may be made from vaginal swabs of females that have aborted within the previous
24 hours, or from the moist fleece of a freshly aborted or stillborn lamb that has not been cleaned by its
mother, or from the abomasal content of the aborted or stillborn lamb. In general, such preparations contain
fewer organisms than placental smears.
In terms of morphology and staining characteristics, C. abortus resembles the rickettsia Coxiella burnetii,
which, in some circumstances, may provoke abortion and which, in humans, causes Q fever. Care must be
taken to differentiate between these two organisms in cases lacking a good history or evidence of
chlamydial-induced placental pathology. Antigenic differences between C. abortus and Coxiella burnetii can
be detected serologically. Fluorescent antibody tests (FATs) using a specific antiserum or monoclonal
antibody may be used for identification of C. abortus in smears.
b)
Antigen detection
Several chlamydial genus-level antigen-detection tests are available commercially. A comparative
assessment of several such assays, on non-ovine material, indicated that those using enzyme-linked
immunosorbent assay (ELISA) methodology were more sensitive than kits employing a FAT (Wood &
Timms, 1992). Under the test conditions used, a kit that detects chlamydial lipopolysaccharide (LPS) was
judged to be the most sensitive of the rapid ELISA-based systems investigated. Though occasionally
yielding false-positive results, particularly with avian faecal samples, the kit also gave satisfactory results
with ovine placental samples (Wilsmore & Davidson, 1991) although it should be noted that it does not
differentiate between C. abortus and other chlamydial species that may contaminate the samples. In
histopathological sections, antigen detection can be performed using commercially available anti-Chlamydia
antibodies directed against LPS or MOMP (major outer membrane protein) (Borel et al., 2006).
c)
DNA
Amplification of chlamydial DNA by polymerase chain reaction (PCR) and real-time PCR provide alternative
approaches for verifying the presence of chlamydiae in biological samples without resorting to culture. PCR
is highly sensitive for this purpose, but has the attendant risk of cross-contamination between samples or
environmental contamination of samples in the field, so appropriate measures must be taken to avoid this
happening (see Guideline 3.2 Biotechnology in the diagnosis of infectious diseases). Another potential
problem is in the production of false negatives resulting from PCR-inhibitory substances in the samples.
Methods for discriminating between amplified DNA sequences originating from C. abortus and C. pecorum
have been described (DeGraves et al., 2003; Everett & Andersen, 1999; Jee et al., 2004; Laroucau et al.,
2001; Thiele et al., 1992). In the last few years, real-time PCR has become the preferred method in
diagnostic laboratories for its rapidity, high throughput and ease of standardisation (Sachse et al., 2009).
Recently, DNA microarray hybridisation assays using the ArrayTube™ platform have been developed and
hold much promise for the direct detection and identification of organisms from clinical samples (Borel et al.,
2008; Sachse et al., 2005). PCR assays in combination with restriction fragment length polymorphism
analysis have been developed with potential to differentiate naturally infected from vaccinated animals
(DIVA) (Laroucau et al., 2010; Wheelhouse et al., 2010).
d)
Tissue sections
Intracellular chlamydial inclusions can be demonstrated by Giemsa staining of thin (≤4 µm) sections taken
from target tissues that have been suitably fixed in fluids such as Bouin or Carnoy. More striking results can
be obtained by immunological staining procedures. The direct immunoperoxidase method (Finlayson et al.,
1985) is rapid and simple, while the method with streptavidin–biotin is more complex (Szeredi & Bacsadi,
2002). Electron microscopy can also be performed using negative contrast, to differentiate chlamydiae from
Coxiella burnetii.
e)
Isolation of the agent
Chlamydophila abortus can be isolated in embryonated chicken eggs or in cell culture, the latter being the
method of choice for isolation of new strains. The causative agent of chlamydiosis is zoonotic and thus
isolation and identification procedures should be carried out under biosafety level 2 conditions.
Tissue samples, such as diseased cotyledons, placental membranes, fetal lung or liver, or vaginal swabs,
that may be subject to any delay before isolation procedures begin, should be maintained in a suitable
transport medium in the interim period. For optimal recovery such samples should be stored frozen,
preferably at –80°C, or otherwise at –20°C. The most satisfactory medium is sucrose/phosphate/glutamate
or SPG medium (sucrose [74.6 g/litre], KH2PO4 [0.512 g/litre], K2HPO4 [1.237 g/litre], L-glutamic acid
[0.721 g/litre]) supplemented with 10% fetal bovine serum, antibiotic (streptomycin and gentamycin are
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suitable, but not penicillin), and a fungal inhibitor (Spencer & Johnson, 1983). A tissue:medium ratio of 1:10
is commonly employed. Alternatively, approximately 1 g of tissue is ground with sterile sand in 8 ml of
transport medium.
Chicken embryos: Test samples are prepared as 10% suspensions in nutrient broth containing streptomycin
(not penicillin) (200 µg/ml); 0.2 ml of suspension is inoculated into the yolk sac of 6–8-day old embryos,
which are then further incubated at 37°C. Infected embryos die between 4 and 13 days after inoculation.
Smears prepared from their vascularised yolk sac membranes reveal large numbers of elementary bodies.
Cell cultures: Chlamydophila abortus of ovine origin can be isolated in a variety of cell types, but McCoy,
Buffalo Green Monkey (BGM) or baby hamster kidney (BHK) cells are most commonly used. For
confirmatory diagnosis, cultured cell monolayers are suspended in growth medium at a concentration of 2 ×
105 cells/ml. Aliquots of 2 ml of the suspension are dispensed into flat-bottomed glass Universal bottles,
each containing a single 16 mm cover-slip. Confluent cover-slip monolayers are achieved after incubation for
24 hours at 37°C. The growth medium is removed and replaced by 2 ml of test inoculum, which is then
centrifuged at 2500 g for 30 minutes on to the cover-slip monolayer to promote infection. After further
incubation for 2–3 days, the cover-slip monolayers are fixed in methanol and stained with Giemsa or
according to the method of Gimenez (Arens & Weingarten, 1981; Gimenez, 1964). After methanol fixation,
infected cultures contain basophilic (Giemsa) or eosinophilic (Gimenez) intracytoplasmic inclusions. Similar
procedures are used in culturing C. abortus for antigen preparation. FAT techniques can also be used and
are equally effective.
Chlamydial activity can be further enhanced by chemical treatment of cultured cells, before or during
infection, to favour chlamydial growth. Various substances that have been described for incorporation into
the infective inoculum to which cover-slip monolayers are exposed include: cycloheximide (0.5 µg/ml) in the
maintenance medium, emetine (1 µg/ml) for 5 minutes before infection, and 5-iodo-2-deoxyuridine
(80 µg/ml) for 3 days prior to infection. Unless preconditioned cells are available, the latter isolation
procedure requires increased time for successful agent isolation.
2.
Serological tests
a)
Complement fixation test
Complement fixation (CF) is the most widely used procedure for detecting infection (sheep and goats are
generally tested within 3 months of abortion or parturition). The test will also detect evidence of vaccination.
Infection is evident principally during active placental infection in the last month of gestation and following
the bacteraemia that often accompanies abortion. Consequently, paired sera collected at the time of
abortion and again at least 3 weeks later may reveal a rising CF antibody titre that will provide a basis for a
retrospective diagnosis. Antigenic cross-reactivity between C. abortus and C. pecorum, as well as with some
Gram-negative bacteria (e.g. Acinetobacter), can give rise to low false-positive CF test results. Thus, titres
less than 1/32 in individual animals should be considered to be nonspecific for C. abortus, although they
could also be due to a low grade infection with C. abortus. Ambiguous results can be investigated further by
western blot analysis using purified elementary bodies (Jones et al., 1997).
Antigen is prepared from heavily infected yolk sac membranes obtained from chicken embryos that have
been inoculated in the same manner as those used to isolate the organism from field material. The
preparation of the antigen should be carried out in a biosafety cabinet with the appropriate biosecurity
precautions to prevent human infection (see chapter 1.1.3). Chopped and ground membranes are
suspended in phosphate buffer, pH 7.6, at the rate of 2 ml per g membrane. After removal of crude debris,
the supernatant fluid is centrifuged at 10,000 g for 1 hour at 4°C, the deposit is resuspended in a small
volume of saline, and a smear of this is examined to ensure a high yield of chlamydiae. The suspension is
held in a boiling water bath for 20 minutes, or is autoclaved, and sodium azide (0.3%) is added as a
preservative. Antigen may also be prepared from cell cultures infected with C. abortus. Infected monolayers
are suspended in phosphate buffer, pH 7.6, and the cells are disrupted by homogenisation or
ultrasonication. Gross debris is removed and subsequent procedures are as for the preparation of antigen
from infected yolk sacs. In either case, CF tests with standardised complement and antisera will establish
the optimal working dilution for each batch of antigen.
b)
Other tests
The serological responses to C. abortus and C. pecorum can be resolved by indirect micro-immunofluorescence, but the procedure is too time-consuming for routine diagnostic purposes. ELISAs developed
independently by several research groups have not been adapted for general diagnostic work, partly
because of difficulties associated with the use of particulate antigens. However, a novel ELISA that
incorporates a stable, solubilised antigen has been used to test experimental and field samples, and has
given results that, though lacking species specificity, have a higher sensitivity than the CF test (Anderson et
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Chapter 2.7.7. — Enzootic abortion of ewes (ovine chlamydiosis)
al., 1995; Jones et al., 1997). Other tests using monoclonal antibody technology in a competitive ELISA
(Salti-Montesanto et al., 1997) and recombinant antigen technology in indirect ELISAs (Longbottom et al.,
2002) have been developed and shown to be more sensitive and specific than the CF test in differentiating
animals infected with C. abortus from those infected with C. pecorum. However, these tests are currently
mainly used as research tools, and have not been developed commercially. A number of commercially
available serological tests have been evaluated and compared with these ‘in-house’ tests with variable
results (Jones et al., 1997; Vretou et al., 2007, Wilson et al., 2009). None of the serological tests that is
available can differentiate vaccination titres from those acquired as a result of natural infection (Borel et al.,
2005).
C. REQUIREMENTS FOR VACCINES
1.
Background
a)
Rationale and intended use of the product
Currently, two types of vaccine (inactivated and attenuated live vaccines) are available commercially, to be
administered intramuscularly or subcutaneously at least 4 weeks before breeding to aid in the prevention of
abortion. A multi-component recombinant vaccine against C. abortus remains a future goal of chlamydial
vaccine research (Longbottom & Livingstone, 2006).
Inactivated vaccines can be prepared from infected yolk sacs or cell cultures (Jones et al., 1995) and
incorporate whole organisms or fractions of them (Tan et al., 1990) using the appropriate biosecurity
precautions to prevent human infection (see chapter 1.1.3). Operator care should be observed in handling
commercial inactivated vaccines that incorporate mineral oil-based adjuvants, as self-injection can result in
severe local inflammation and tissue necrosis. The commercial live, attenuated vaccine is a chemically
induced temperature-sensitive mutant strain of the organism that grows at 35°C but not at 39.5°C, the body
temperature of sheep (Rodolakis, 1986). This vaccine is supplied lyophilised and must be reconstituted in
diluent immediately before administration. Operator care should be observed in handling and administering
this live vaccine, particularly by immunocompromised individuals and pregnant women. Importantly, the live
vaccine must not be given to animals being treated with antibiotics, particularly tetracyclines. Inactivated
vaccines are safe for administration during pregnancy, whereas live vaccines cannot be used in pregnant
animals.
Both types of vaccine have a role to play in controlling disease, but neither confers absolute protection
against challenge or completely reduces the shedding of infective organisms. However, vaccinates exposed
to infection do experience significantly lower abortion rates and reduced excretion of chlamydiae for at least
two to three lambings after vaccination. It has been claimed that the live vaccine could be an aid to
eradication of disease (Nietfeld, 2001). In addition, the live vaccine strain 1B has been detected in the
placentas of vaccinated animals that have aborted as a result of OEA, suggesting a possible role for the
vaccine in causing disease (Wheelhouse et al., 2010), but despite this the use of live vaccine remains the
most effective method of protecting from the disease (Stuen & Longbottom, 2011).
Vaccine stored under refrigeration (5±3°C) should remain stable for at least 1 year. No firm data are
available, but revaccination is recommended every 1–3 years, according to the exposure risk.
2.
Outline of production and minimum requirements for conventional vaccines
a)
Characteristics of the seed
i)
Biological characteristics
One or more ovine abortion isolates that consistently grow productively in the chosen substrate are
suitable, and an early passage of the seed stock can be established. Alternatively, an isolate that has
been adapted to the chicken embryo by multiple passage (>100) can be used. This permits more of the
embryo to be used for vaccine production. Although adaptation to the embryo may diminish the
isolate’s virulence for sheep, there is no evidence that such change reduces its protective efficacy as
an inactivated vaccine.
ii)
Quality criteria (sterility, purity, freedom from extraneous agents)
Before inoculation of large numbers of embryos or cell cultures, the viability and freedom from
contamination of seed stock should be verified. It may be convenient to collect the total harvest in
separate manageable lots. In this case, the infectivity of an aliquot of each lot should be separately
titrated to ensure that each matches the requirements (see below). Store under refrigerated conditions.
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Chapter 2.7.7. — Enzootic abortion of ewes (ovine chlamydiosis)
b)
Method of manufacture
i)
Procedure
For low passage isolates, the procedures described for the preparation of CF antigen are suitably
adapted and amplified for bulk production. Once the final harvest suspension is obtained, an aliquot is
removed for titration of its infectivity. The bulk is treated with formalin to a final concentration of 0.4%,
and stored until sterility tests confirm complete inactivation.
ii)
Requirements for substrates and media
The inactivated harvest is centrifuged and resuspended in phosphate buffered saline containing 0.2%
formalin to a volume representing a preinactivation infectivity titre of approximately 108 infectious
units/ml. Usually, the aqueous suspension is blended with an oil adjuvant, either directly or after
precipitation by potassium alum (AlK[SO4]2.12 H2O). A preservative, such as 0.01% thiomersal, may
also be added.
iii)
In-process controls
The main requirements are to ensure adequate growth of C. abortus, avoidance of extraneous infection
of the culture substrate, completeness of inactivation and biohazard awareness by process workers.
iv)
Final product batch tests
Each separate batch of manufactured vaccine should be tested for sterility, safety and potency.
Sterility and purity
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
Safety
Subcutaneous inoculation into two or more seronegative sheep of twice the standard dose (usually
1.0 ml) of manufactured vaccine should elicit no systemic reaction, but oil-adjuvant vaccines can cause
a nonharmful swelling at the inoculation site.
Batch potency
At present, potency is judged by the occurrence of a serological response in previously unvaccinated
sheep given 1 ml of vaccine subcutaneously. Blood samples taken before and 28 days after
vaccination are compared. Ultimately, potency has to be judged against experimental challenge or field
performance, but no in-vitro correlation of protective efficacy has yet been established.
c)
Requirements for authorisation
i)
Safety requirements
See chapter 1.1.6 Principles of veterinary vaccine production.
ii)
Efficacy requirements
See chapter 1.1.6 .
iii)
Stability
See chapter 1.1.6.
3.
Vaccines based on biotechnology
a)
Vaccines available and their advantages
No biotechnology-based vaccines are currently in use for this disease.
REFERENCES
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Blackwell Scientific Ltd., Oxford, UK, 105-112.
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Chapter 2.7.7. — Enzootic abortion of ewes (ovine chlamydiosis)
ANDERSON I.E., HERRING A.J., JONES G.E., LOW J.C. & GREIG A. (1995). Development and evaluation of an indirect
ELISA to detect antibodies to abortion strains of Chlamydia psittaci in sheep sera. Vet. Microbiol., 43, 1–12.
ARENS M. & WEINGARTEN M. (1981). Vergleichende Untersuchungen an Buffalo Green monkey (BGM) Zellen und
Mausen zur Isolierung von Chlamydia psittaci aus Kot und Organproben von Vogeln. Zentralbl. Veterinarmed [B],
28, 301–309.
BOREL N., KEMPF E., HOTZEL H., SCHUBERT E., TORGERSON P., SLICKERS P., EHRICHT R., TASARA T., POSPISCHIL A. &
SACHSE K. (2008). Direct identification of chlamydiae from clinical samples using a DNA microarray assay – a
validation study. Mol. Cell. Probes, 22, 55–64
BOREL N., SACHSE K., RASSBACH A., BRUCKNER L., VRETOU E., PSARROU E. & POSPISCHIL A. (2005). Ovine enzootic
abortion (OEA): antibody response in vaccinated sheep compared to naturally infected sheep. Vet. Res.
Commun., Suppl 1, 151–156.
BOREL N., THOMA R., SPAENI P., WEILENMANN R., TEANKUM K., BRUGNERA E., ZIMMERMANN D.R., VAUGHAN L. &
POSPISCHIL A. (2006). Chlamydia-related abortions in cattle from Graubunden, Switzerland. Vet. Pathol., 43, 702–
708.
BUXTON D., ANDERSON I.E., LONGBOTTOM D., LIVINGSTONE M., WATTEGADERA S. & ENTRICAN G. (2002). Ovine
chlamydial abortion: characterization of the inflammatory immune response in placental tissues. J. Comp. Pathol.,
127, 133–141.
DEGRAVES F.J., GAO D., HEHNEN H.-R., SCHLAPP T. & KALTENBOECK B. (2003). Quantitative detection of Chlamydia
psittaci and C. Pecorumby high-sensitivity real-time PCR reveals high prevalence of vaginal infection in cattle. J.
Clin. Microbiol., 41, 1726–1729.
ENTRICAN G. (2002). Immune regulation during pregnancy and host-pathogen interactions in infectious abortion. J.
Comp. Pathol., 126, 79–94.
EVERETT K.D. & ANDERSEN A.A. (1999). Identification of nine species of the Chlamydiaceae using PCR RFLP. Int.
J. Syst. Bacteriol., 49, 803–813.
EVERETT K.D.E., BUSH R.M. & ANDERSEN A.A. (1999). Emended description of the order Chlamydiales, proposal of
Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus of the family
Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms.
Int. J. System. Bact., 49, 415–440.
FINLAYSON J., BUXTON D., ANDERSON I.E. & DONALD K.M. (1985). Direct immunoperoxidase method for
demonstrating Chlamydia psittaci in tissue sections. J. Clin. Pathol., 38, 712–714.
GIMENEZ D.F. (1964). Staining rickettsiae in yolk-sac cultures. Stain Technol., 39, 135–140.
GUTIERREZ J., WILLIAMS E.J., O’DONOVAN J., BRADY C., PROCTOR A.F., MARQUES P.X., WORRALL S., NALLY J.E.,
MCELROY M., BASSETT H.F., SAMMIN D.J. & MARKEY B.K.(2011).Monitoring clinical outcomes, pathological changes
and shedding of Chlamydophila abortus following experimental challenge of periparturient ewes utilizing the
natural route of infection. Vet. Microbiol., 147 (1–2), 119–126.
JEE J., DEGRAVES F.J., KIM T. & KALTENBOECK B. (2004). High prevalence of natural Chlamydophila species
infection in calves. J. Clin. Microbiol., 42, 5664–5672.
JONES G.E., JONES K.A., MACHELL J., BREBNER J., ANDERSON I.E. & HOW S. (1995). Efficacy trials with tissue-culture
grown, inactivated vaccines against chlamydial abortion in sheep. Vaccine, 13, 715–723.
JONES G.E., LOW J.C., MACHELL J. & ARMSTRONG K. (1997). Comparison of five tests for the detection of antibodies
against chlamydial (enzootic) abortion of ewes. Vet. Rec., 141, 164–168.
LAROUCAU K., SOURIAU A. & RODOLAKIS A. (2001). Improved sensitivity of PCR for Chlamydophila using pmp
genes. Vet. Microbiol., 82 155–164.
LAROUCAU K., VORIMORE F., SACHSE K., VRETOU E., SIARKOU V.I., WILLEMS H., MAGNINO S., RODOLAKIS A. & BAVOIL
P.M. (2010). Differential identification of Chlamydophila abortus live vaccine strain 1B and C. abortus field isolates
by PCR-RFLP. Vaccine, 28, 5653–5656.
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LIVINGSTONE M., WHEELHOUSE N., MALEY S.W. & LONGBOTTOM D. (2009). Molecular detection of Chlamydophila
abortus in post-abortion sheep at oestrus and subsequent lambing. Vet. Microbiol., 135 (1–2), 134–141.
LONGBOTTOM D. & COULTER L.J. (2003). Animal chlamydioses and zoonotic implications. J. Comp. Pathol., 128,
217–244.
LONGBOTTOM D., FAIRLEY S., CHAPMAN S., PSARROU E., VRETOU E. & LIVINGSTONE M. (2002). Serological diagnosis of
ovine enzootic abortion by enzyme-linked immunosorbent assay with a recombinant protein fragment of the
polymorphic outer membrane protein POMP90 of Chlamydophila abortus. J. Clin. Microbiol. 40, 4235–4243.
LONGBOTTOM D. & LIVINGSTONE M. (2006). Vaccination against chlamydial infections of man and animals. Vet. J.,
171, 263–275.
NIETFELD J.C. (2001). Chlamydial infections in small ruminants. Update on Small Ruminant Medicine, 17, 2.
PAPP J.R., SHEWEN P.E. & GARTLEY (1994). Abortion and subsequent excretion of chlamydiae from the
reproductive tract of sheep. Infect. Immun., 62, 3786–3792.
RODOLAKIS A. (1986). Use of a live temperature-sensitive vaccine in experimental and natural infections. In:
Chlamydial Diseases of Ruminants, Aitken I.D., ed. Commission of the European Communities, Luxembourg, 71–
77.
SACHSE K., HOTZEL H., SLICKERS P., ELLINGER T. & EHRICHT R. (2005). DNA microarray-based detection and
identification of Chlamydia and Chlamydophila spp. Mol. Cell Probes, 19, 41–50.
SACHSE K., VRETOU E., LIVINGSTONE M., BOREL N., POSPISCHIL A. & LONGBOTTOM D. (2009). Recent developments in
the laboratory diagnosis of chlamydial infections (Review). Vet. Microbiol., 135, 2–21.
SALTI-MONTESANTO V., TSOLI E., PAPAVASSILIOU P., PSARROU E., MARKEY B.M., JONES G.E. & VRETOU E. (1997).
Diagnosis of ovine enzootic abortion, using a competitive ELISA based on monoclonal antibodies against variable
segments 1 and 2 of the major outer membrane protein of Chlamydia psittaci serotype 1. Am. J. Vet. Res., 58,
228–235.
SILLIS M. & LONGBOTTOM D. (2011). Chlamydiosis. In: Oxford Textbook of Zoonoses, Biology, Clinical Practice and
Public Health Control, Palmer S.R., Lord Soulsby, Torgerson P.R. & Brown D.W.G., eds. Oxford University Press,
Oxford, UK, 146–157.
SPENCER W.N. & JOHNSON F.W.A. (1983). Simple transport medium for the isolation of Chlamydia psittaci from
clinical material. Vet. Rec., 113, 535–536.
STAMP J.T., MCEWEN A.D., WATT J.A.A. & NISBET D.I. (1950). Enzootic abortion in ewes. I. Transmission of the
disease. Vet. Rec., 62, 251–254.
STUEN S. & LONGBOTTOM D. (2011). Treatment and control of Chlamydial and Rickettsial infections in sheep and
goats. Vet. Clin. Food Anim., 27, 213–233.
SZEREDI L. & BACSADI A. (2002). Detection of Chlamydophila (Chlamydia) abortus and Toxoplasma gondii in
smears from cases of ovine and caprine abortion by the streptavidin-biotin method. J. Comp. Pathol., 127, 257–
263.
TAN T.W., HERRING A.J., ANDERSON I.E. & JONES G.E. (1990). Protection of sheep against Chlamydia psittaci
infection with a subcellular vaccine containing the major outer membrane protein. Infect. Immun., 58, 3101–3108.
THIELE D., WITTENBRINK M.M., FISCHER D. & KRAUSS H. (1992). Evaluation of the polymerase chain reaction (PCR)
for detection of Chlamydia psittaci in abortion material from ewes. Zentralbl. Bakteriol., 277, 446–453.
VRETOU E., RADOUANI F., PSARROU, E., KRITIKOS I., XYLOURI E. & MANGANA O. (2007). Evaluation of two commercial
assays for the detection of Chlamydophila abortus antibodies. Vet. Microbiol., 123, 153–161.
WHEELHOUSE N., AITCHISON K., LAROUCAU K., THOMSON J. & LONGBOTTOM D. (2010). Evidence of Chlamydophila
abortus vaccine strain 1B as a possible cause of ovine enzootic abortion. Vaccine, 28 (35), 5657–5663.
WILSMORE A.J. & DAVIDSON I. (1991). ‘Clearview’ rapid test compared with other methods to diagnose chlamydial
infection. Vet. Rec., 128, 503–504.
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WILSON K., LIVINGSTONE M. & LONGBOTTOM D. (2009). Comparative evaluation of eight serological assays for
diagnosing Chlamydophila abortus infection in sheep. Vet. Microbiol. 135, 38-45.
WOOD M.M. & TIMMS P. (1992). Comparison of nine antigen detection kits for diagnosis of urogenital infections due
to Chlamydia psittaci in koalas. J. Clin. Microbiol., 30, 3200–3205.
*
* *
NB: There are OIE Reference Laboratories for Enzootic abortion of ewes
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for Enzootic abortion of ewes
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CHAPTER 2.7.8.
NAIROBI SHEEP DISEASE
See Chapter 2.9.1. Bunyaviral diseases of animals (excluding Rift Valley fever)
*
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2009
CHAPTER 2.7.9.
OVINE EPIDIDYMITIS
(Brucella ovis)
SUMMARY
Brucella ovis produces a clinical or subclinical disease in sheep that is characterised by genital
lesions in rams, and placentitis in ewes. Accordingly, the main consequences of the disease are
reduced fertility in rams, infrequent abortions in ewes, and an increased perinatal mortality. The
disease has been reported in Latin American, North American and European countries as well as
Australia, New Zealand and South Africa, but probably occurs in most sheep-raising countries.
Identification of the agent: The existence of clinical lesions (unilateral or, occasionally, bilateral
epididymitis) in rams may be indicative of the existence of infection, but laboratory examinations are
necessary to confirm the disease. Laboratory confirmation may be based on direct or indirect
methods. Direct diagnosis is made by means of bacteriological isolation of B. ovis from semen
samples or tissues of rams, or vaginal discharges and milk of ewes, on adequate selective media.
Molecular biological methods have being developed that could be used for complementary
identification based on specific genomic sequences. The polymerase chain reaction (PCR)
methods provide additional means of detection. However, indirect diagnosis based on serological
tests is preferred for routine diagnosis.
Serological tests: The complement fixation test (CFT), agar gel immunodiffusion (AGID) test and
indirect enzyme-Iinked immunosorbent assay (I-ELISA) using soluble surface antigens obtained
from B. ovis, can be used. Some I-ELISAs using recombinant proteins and monoclonal antibodies
are being tested in field trials. The sensitivities of the AGID test and ELISA are similar and
sometimes the I-ELISA has higher sensitivity than the CFT. A combination of the AGID test and IELISA seems to give the best results in terms of sensitivity. However, with regard to simplicity and
cost, the AGID test is the most practicable test for diagnosis of B. ovis. However, because of the
lack of standardised methods recognised at the international level for I-ELISA and AGID, the
prescribed test for international trade remains the CFT.
Requirements for vaccines and diagnostic biologicals: Seed cultures for antigen or vaccine
production should be obtained from internationally recognised laboratories. A single standard dose
(109 colony-forming units) of the live B. melitensis Rev.1 vaccine, administered subcutaneously or
conjunctivally, can be used safely and effectively in rams, for the prevention of B. ovis infection.
This vaccine strain should meet minimal quality standards: adequate concentration, absence of
dissociation, adequate residual virulence and immunogenicity and free of extraneous agents (see
Chapter 2.7.2 Caprine and ovine brucellosis [excluding B. ovis]).
A. INTRODUCTION
Brucella ovis causes a genital infection of ovine livestock manifested by epididymitis, infrequent abortions, and
increased lamb mortality. Passive venereal transmission via the ewe appears to be a frequent route of infection,
but ram-to-ram transmission is also common1 (Blasco, 1990). Infected ewes may excrete B. ovis in vaginal
1
Under the semi-extensive production systems (most common in European Mediterranean countries) rams are usually
housed together. Direct ram-to-ram transmission during non-breeding periods is quite frequent and has been suggested to take
place by several routes, including the rectal mucosa. Most ram-to-ram infections, however, are produced through the oral route.
Housed rams establish hierarchies (head-to-head combats), and it is frequent that ‘dominated’ rams, after being ‘mated’ by the
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
discharges and milk and, accordingly, ewe-to-ram and lactating ewe-to-lamb transmission could also be
determinant mechanisms of infection. Accordingly, the ewes are as relevant as rams in the epidemiology of
infection, and control or eradication of B. ovis is feasible only if females are included in the corresponding
programme.
The demonstration of the existence of genital lesions (unilateral or, occasionally, bilateral epididymitis) by
palpating the testicles of rams may be indicative of the presence of this infection in a given flock. However, this
clinical diagnosis is not sensitive enough because only about 50% of rams infected with B. ovis present
epididymitis (Blasco, 1990). Moreover, the clinical diagnosis is extremely nonspecific due to the existence of
many other bacteria causing clinical epididymitis. The most frequently reported isolates causing epididymitis in
rams include Actinobacillus seminis, A. actinomycetemcomitans, Histophilus ovis, Haemophilus spp.,
Corynebacterium pseudotuberculosis ovis, B. melitensis and Chlamydophila abortus (formerly Chlamydia psittaci)
(Bulgin & Anderson, 1983; Burgess & McDowell, 1981; De Long et al., 1979; Ekdahl M.O., Money & Martin, 1968;
Livingstone & Hardy, 1964; Rodolakis & Bernard, 1977; Webb, 1983; Williamson & Nairn, 1980). It must be
emphasised that many palpable epididymal lesions in rams are sterile, trauma-induced spermatic granulomas.
Although cattle, goats and deer have been proved susceptible to B. ovis in artificial transmission experiments,
natural cases have been reported only in deer (Ridler, 2001). To date, no human cases have been reported, and
B. ovis is considered to be non-zoonotic. However, in areas where B. melitensis infection co-exists with B. ovis,
special care is required when handling samples, which should be transported to the laboratory in leak-proof
containers (for further details see Chapter 2.4.3 Bovine brucellosis).
The classification, microbiological and serological properties of the genus Brucella and related species and
biovars are given in the chapter 2.4.3.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
a)
Collection of samples
The most valuable samples for the isolation of B. ovis from live animals are semen, vaginal swabs and milk.
For the collection of vaginal swabs and milk, see the instructions given in Chapter 2.7.2 Caprine and ovine
brucellosis (excluding B. ovis). Semen (genital fluids) can be collected easily in swabs taken from the
preputial cavity after electro-ejaculation. lf an electro-ejaculator is not available, swabs can be taken from the
vagina of brucellosis-free ewes immediately after natural mating.
For the isolation of B. ovis after necropsy, the preferred organs in terms of probability of isolation are the
epididymides, seminal vesicles, ampullae, and inguinal lymph nodes in rams, and the uterus, iliac and supramammary lymph nodes in ewes. However, to obtain maximum sensitivity, a complete search that includes
other organs and lymph nodes (spleen, cranial, scapular, prefemoral and testicular lymph nodes) should be
performed. Dead lambs and placentas should also be examined. The preferred culture sites in aborted or
stillborn lambs are abomasal content and lung.
Samples for culture should be refrigerated and transported to the laboratory to be cultured as soon as
possible after collection. The organism remains viable for at least 72 hours at room temperature and survival
is enhanced at 4°C or, preferably, by freezing the tissue samples.
b)
Staining methods
Semen or vaginal smears can be examined following staining by Stamp’s method (Alton et al., 1988; Corbel
et al., 1978) (see chapter 2.7.2), and characteristic coccobacilli should be demonstrated in many infected
animals (Webb et al., 1980). Examination of Stamp-stained smears of suspect tissues (ram genital tract,
inguinal lymph nodes, placentas, and abomasal content and lung of fetuses) may also allow a rapid
presumptive diagnosis.
However, other bacteria with similar morphology or staining characteristics (B. melitensis, Coxiella burnetii,
and Chlamydophila abortus) can also be present in such samples, making the diagnosis difficult for
inexperienced personnel. Microscopy results should always be confirmed by culture of the microorganism.
dominant rams, lick the prepuce of these dominant rams as an act of submission. If these dominant rams are infected, the
probability of having B. ovis in the prepuce (excretion in the semen) is very high.
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
c)
Culture
The best direct method of diagnosis is bacteriological isolation on adequate culture media. Semen, vaginal
swabs, or milk samples can be smeared directly on to plates with adequate culture media and incubated at
37°C in an atmosphere of 5–10% CO2. Tissues should be macerated and ground in a small amount of
sterile saline or phosphate buffered saline (PBS) with a stomacher or blender, before plating.
Growth normally appears after 3–4 days, but cultures should not be discarded as negative until 7 days have
elapsed. Colonies of B. ovis become visible (0.5–2.5 mm) after 3–4 days of incubation, and are rough
phase, round, shiny and convex.
Brucella ovis can be isolated in nonselective media, such as blood agar base enriched with 10% sterile
ovine or bovine sera, or in blood agar medium with 5–10% sterile ovine blood. However, the inoculum
frequently contains other bacteria, which often overgrow B. ovis. Accordingly, the use of selective media
may be preferred. Various B. ovis selective media have been described. The modified Thayer–Martin’s
medium (Brown et al., 1971; Marin et al., 1996) is recommended. Briefly, it can be prepared with GC
medium base (38 g/litre; Biolife Laboratories, Milan, Italy) supplemented with haemoglobin (10 g/litre; Difco)
and colistin methane-sulphonate (7.5 mg/litre), vancomycin (3 mg/litre), nitrofurantoin (10 mg/litre), nystatin
(100,000 International Units [IU]/litre = 17.7 mg) and amphotericin B (2.5 mg/litre) (all products from Sigma
Chemical, St Louis, United States of America [USA]). Working solutions are prepared as follows:
Solution A: Add 500 ml of distilled water to the GC medium base, heat the paste carefully while stirring
continuously and autoclave at 120°C for 20 minutes.
Solution B: Suspend the haemoglobin in 500 ml of distilled water, adding the water slowly to avoid lumps.
Once dissolved, add a magnetic stirrer and autoclave at 120°C for 20 minutes.
Antibiotic solution (prepared daily): colistin, nystatin and vancomycin are suspended in a mixture of
methanol/water (1/1); nitrofurantoin is suspended in 1 ml of a 0.1 M NaOH sterile solution. For amphotericin
B, it is recommended to prepare a stock solution of 10 mg/ml amphotericin B with 10 mg dissolved first in
1 ml sterile dimethyl sulphoxide (C2H6OS, for analysis; ACS) and then added to 9 ml of PBS (10 mM,
pH 7.2). Any stock solution remaining can be stored some days at 4°C. All antibiotic solutions must be
filtrated through 0.22 µm filters before addition to the culture medium.
Once autoclaved, stabilise the temperature (45–50°C) of both solutions A and B with continuous stirring. Mix
both solutions (adding A to B), avoiding bubble formation. Add the antibiotic solutions while stirring
continuously and carefully. Dispense into sterile plates.
Once prepared, the plates should not be stored for long periods, and freshly prepared medium is always
recommended. This medium is also suitable for the isolation of B. melitensis (see chapter 2.7.2).
All culture media should be subjected to quality control with the reference strain, to show that it supports
growth.
Another suitable, but less effective, antibiotic combination is: vancomycin (3 mg/litre); colistin (7.5 mg/litre);
nystatin (12,500 IU/litre); and nitrofurantoin (10 mg/litre).
The Farrell’s medium described for the culture of smooth brucellae is not appropriate for the culture of
B. ovis as it does not grow on this medium.
d)
Identification and typing
Brucella ovis colonies are not haemolytic. They are circular, convex, have unbroken edges, are always of
the rough type when examined by oblique illumination, and test positive in the acriflavine test (Alton et al.,
1988; Corbel et al., 1978). For growth, B. ovis needs an atmosphere of 5–10% CO2. It lacks urease activity,
fails to reduce nitrate to nitrite, is catalase positive and oxidase negative. It does not produce H2S and,
although it does not grow in the presence of methyl violet, it usually grows in the presence of standard
concentrations of basic fuchsin and thionin. The cultures are not lysed by Brucella-phages of the Tbilissi
(Tb), Weybridge (Wb) and Izatnagar (Iz) groups at the routine test dilution (RTD) or 104 RTD, while they are
lysed by phage R/C (Alton et al., 1988; 7). Most laboratories are not equipped for a complete identification,
and a practical schedule for presumptive identification is needed. Most B. ovis isolates can be correctly
identified on the basis of growth characteristics, direct observation using obliquely reflected light, Gram or
Stamp’s staining, catalase, oxidase, urease and acriflavine tests. However, definitive identification should be
carried out by reference laboratories with experience in identification and typing of Brucella.
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
The polymerase chain reaction (PCR) and other recently developed molecular methods provide additional
means of detection and identification of Brucella sp. (see chapter 2.4.3).
2.
Serological tests
The most efficient and widely used tests are the complement fixation test (CFT), the double agar gel
immunodiffusion (AGID) test and the indirect enzyme-Iinked immunosorbent assay (I-ELISA). Several countries
have adopted various standard diagnostic techniques for B. ovis, but the only test prescribed by the OIE and the
European Union (EU) for international trade is the CFT. However, it has been demonstrated that the AGID test
shows similar sensitivity to the CFT, and it is a simpler test to perform. Although standardisation is lacking,
numerous independent studies have shown that the I-ELISA is more sensitive and specific than either the CFT or
AGID test, and with further validation and standardisation studies, the I-ELISA could become a suitable candidate
for future designation as a prescribed test for B. ovis diagnosis.
The International Standard anti-Brucella ovis Serum (International Standard 19852) is the one against which all
other standards are compared and calibrated. This reference standard is available to national reference
laboratories and should be used to establish secondary or national standards against which working standards
can be prepared and used in the diagnostic laboratory for daily routine use.

Antigens
When rough Brucella cells are heat-extracted with saline (hot-saline method, HS), they yield water-soluble
antigenic extracts, the major component of which precipitates with sera to rough Brucella (Diaz & Bosseray,
1973; Myers et al., 1972). For this reason, the HS extracts have been referred to as the ‘rough-specific
antigen’ or, when obtained from B. ovis, as the ‘B.-ovis-specific antigen’. However, the chemical
characterisation of the HS extracts from B. ovis has shown that they are enriched in rough
lipopolysaccharide (R-LPS), group 3 outer membrane proteins and other outer membrane components
(Riezu-Boj et al., 1986). Thus, HS extracts contain LPS determinants specific for B. ovis, but also additional
antigenic components, some of them shared with rough and smooth B. melitensis and other Brucella
(Santos et al., 1984). Such components account for the cross-reactivity that is sometimes observed with the
HS method and sera of sheep infected with B. melitensis or vaccinated with Rev.1 (Riezu-Boj et al., 1986).
The HS extract, due to its water solubility and high content of relevant cell-surface epitopes, is the best
diagnostic antigen and has been widely used for the serological diagnosis of B. ovis infection.
Brucella ovis REO 198, a CO2- and serum-independent strain, is recommended as a source of the HS
antigens to be used in serological tests3. Solid media described in Section B.1.c. are satisfactory for the
growth of B ovis REO 198. HS antigen is prepared as follows:
i)
Exponentially grow a suitable strain of B ovis, preferably aerobic and nonserum dependent, e.g. REO
198, in one of the following ways: for 48 hours in trypticase–soy broth flasks in an orbital incubator at
37°C and 150 rpm; or in Roux bottles of trypticase–soy agar, or other suitable medium, with 5% serum
added (not necessary when using the REO 198 strain); or in a batch-type fermenter as described for
B. abortus, but with the addition of 5% serum to the medium (not necessary when using the REO
198 strain).
ii)
Cells are resuspended in 0.85% saline or PBS, then washed twice in 0.85% saline (12 g of dried cells
or 30 g of wet packed cells in 150 ml).
iii)
The cell suspension is then autoclaved at 120°C for 15–30 minutes.
iv)
After cooling, the suspension is centrifuged (15,000 g, 4°C, 15 minutes) and the supernatant fluid is
filtered and dialysed against distilled water using 100 times the volume of the suspension, at 4°C; the
water should be changed three times over a minimum of 2 days.
v)
The dialysed fluid can be ultracentrifuged (100,000 g, 4°C, 6–8 hours), and the sediment is
resuspended in a small amount of distilled water and freeze-dried. When produced to be used in the
CFT, the addition of control process serum replacement II (CPSRII) prior to freeze-drying may assist in
stability and anti-complementary activity.
HS is then resuspended either in distilled water (for use in the AGID test), veronal buffered saline (for use in
the CFT), or carbonate/bicarbonate buffer or PBS (for use in the I-ELISA) and titrated against a set of
adequate positive and negative sera.
2
3
Obtainable from the OIE Reference Laboratory for Brucellosis at AHVLA Weybridge, Addlestone, Surrey KT15 3NB,
United Kingdom.
Obtainable from the OIE Reference Laboratory for Brucellosis at Anses Maisons-Alfort, 94706, France.
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
If it is to be used in the AGID test, the resuspended HS could be kept at 4°C adding 0.5% phenol as
preservative. Freezing and thawing should be avoided (Diaz & Bosseray, 1973). The CFT antigen should be
standardised against the International anti-B. ovis Standard Serum to give 50% fixation at a 1/100 serum
dilution.
a)
Complement fixation test (the prescribed test for international trade)
There is no standardised method for the CFT, but the test is most conveniently carried out using the
microtitration method. Some evidence shows that cold fixation is more sensitive than warm fixation (Burgess
& Norris, 1982; Ris et al., 1984; Searson, 1982), but that it is less specific. Anticomplementary reactions,
common with sheep serum, are, however, more frequent with cold fixation.
Several methods have been proposed for the CFT using different concentrations of fresh sheep red blood
cells (SRBCs) (a 2–3% suspension is usually recommended) sensitised with an equal volume of rabbit antiSRBC serum diluted to contain several times (usually from two to five times) the minimum concentration
required to produce 100% lysis of SRBCs in the presence of a titrated solution of guinea-pig complement.
The latter is independently titrated (in the presence or absence of antigen according to the method) to
determine the amount of complement required to produce either 50% or 100% lysis of sensitised SRBCs in
a unit volume of a standardised suspension; these are defined as the 50% or 100% haemolytic unit of
complement (C’H50 or C’H100), respectively. It is generally recommended to titrate the complement before
each set of tests, a macromethod being preferred for an optimal determination of C’H50. Usually, 1.25–
2 C’H100 or 5–6 C’H50 are used in the test.
Barbital (veronal) buffered saline (VBS) is the standard diluent for the CFT. This is prepared from tablets
available commercially, otherwise it may be prepared according to the formula described elsewhere (see
Chapter 2.4.3 Bovine brucellosis). The test sera should be inactivated for 30 minutes in a water bath at 60–
63°C, and then diluted (doubling dilutions) in VBS. The stock solution of HS antigen (2.5–20 mg/ml in VBS)
is diluted in VBS as previously determined by titration (checkerboard titration). Usually, only one serum
dilution is tested (generally 1/10).
Using standard 96-well microtitre plates with round (U) bottom, the technique is usually performed as
follows:
i)
Volumes of 25 µl of diluted inactivated test serum are placed in the well of the first and second rows.
Volumes of 25 µl of CFT buffer are added to all wells except those of the first row. Serial doubling
dilutions are then made by transferring 25 µl volumes of serum from the second row onwards.
ii)
Volumes of 25 µl of antigen, diluted to working strength, are added to each well except wells in the first
row.
iii)
Volumes of 25 µl of complement, diluted to the number of units required, are added to each well.
iv)
Control wells containing diluent only, complement + diluent, antigen + complement + diluent, are set up
to contain 75 µl total volume in each case. A control serum that gives a minimum positive reaction
should be tested in each set of tests to verify the sensitivity of test conditions.
v)
The plates are incubated at 37°C for 30 minutes or at 4°C overnight, and a volume (25 or 50 µl
according to the techniques) of sensitised SRBCs is added to each well. The plates are reincubated at
37°C for 30 minutes.
vi)
The results are read after the plates have been centrifuged at 1000 g for 10 minutes at 4°C or left to
stand at 4°C for 2–3 hours to allow unlysed cells to settle. The degree of haemolysis is compared with
standards corresponding to 0, 25, 50, 75 and 100% lysis. The titre of the serum under test is the
highest dilution in which there is 50% or less haemolysis.

Standardisation of the results of the complement fixation test
There is a unit system that is based on the International Standard for anti-Brucella ovis Serum
(International Standard 1985 [see footnote 2]). This serum contains 1000 IU/ml. lf this serum is tested
in a given method and gives a titre of, for example 100, then the factor for an unknown serum tested by
that method can be found from the formula: 1000/100 × titre of test serum = number of ICFTU
(International CFT units) of antibody in the test serum per ml. It is recommended that any country using
the CFT on a national scale should obtain agreement among the different laboratories performing the
test by the same method, to allow the same level of sensitivity and specificity to be obtained against an
adequate panel of sera from B. ovis culture positive and Brucella-free sheep. Results should always be
expressed in ICFTU, calculated in relation to those obtained in a parallel titration with a standard
serum, which itself may be calibrated against the International Standard.
1022
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
Interpretation of the results: sera giving a titre equivalent to 50 ICFTU/ml or more are considered to be
positive in the EU.
b)
Agar gel immunodiffusion test
The AGID test (Blasco, 1990) uses the following reagents: Good grade Noble agar or agarose, sodium
chloride (NaCI), and borate buffer (prepared with boric acid [12.4 g]; potassium chloride [14.5 g]; distilled
water [1600 ml]; adjusted to pH 8.3 with 0.2 M NaOH solution and made up to 2000 ml with distilled water).
To prepare the gels, dissolve 1 g of agarose (or Noble agar), 10 g of NaCI and 100 ml of borate buffer by
boiling while stirring continuously. On a flat surface, cover clean glass slides with the necessary amount of
molten gel to form a bed of 2.5 mm depth (3.5 ml approximately for standard microslides). After the gel has
solidified (15–20 minutes), wells are cut in it using a gel puncher. The wells should be 3 mm in diameter and
3 mm apart, and should be arranged in a hexagonal pattern around a central well that is also 3 mm in
diameter. The test can be adapted to Petri dishes and other patterns.
Sera to be examined are placed in alternate wells separated by a control positive serum (infection proved by
bacteriology), with the antigen at its optimum concentration in the central well. The results are read after
incubation for 24 and 48 hours at room temperature in a humid chamber. A positive reaction is a clearly
defined precipitin line between the central well and the wells of the test sera that gives total or partial identity
with that of the positive controls. Precipitin lines not giving total identity may also appear and correspond to
minor antigenic components of HS extracts (antibodies to these components can also be common in
infections due to B. melitensis). These reactions should also be considered to be positive. Before a definitive
reading, it is important to wash the slides for 1 hour in a 5% sodium citrate water solution to clean unspecific
precipitin lines.
The HS (2.5–20 mg/ml) diluted in distilled water (optionally containing 0.5% phenol as a preservative) is the
antigen used in the AGID test (the preserved antigen can be stored at 4°C for at least 1 month). Dilutions of
antigen are tested with a panel of 20–30 sera from rams naturally infected with B. ovis and with a panel of
Brucella-free sheep. The optimum concentration of antigen is that giving the clearest precipitation line with
all the sera from B.-ovis-infected rams being negative with the sera from Brucella-free sheep.
c)
Enzyme-linked immunosorbent assay (the alternative test for international trade)
Several variations of this assay have been proposed. The assay described here is an indirect I-ELISA using
ABTS (2,2’-azino-bis-[3-ethylbenzothiazoline-6-sulphonic acid]) as chromogen, but other procedures are
also suitable. Tests are performed on 96-weIl flat-bottomed ELISA plates. Reagent and serum dilutions are
made in PBS, pH 7.2, with the addition of 0.05% Tween 20 (PBST). Antigen dilutions are made in a
carbonate/bicarbonate buffer, pH 9.6, or, alternatively, in PBS, pH 7.2. Plates are washed after antigen
coating and between incubations, where appropriate, usually with PBST. The antigen (HS) and conjugate
are checkerboard titrated, and dilutions are selected to give the best discriminating ratio between negative
and positive standard sera. Secondary antibodies (anti-ovine IgG [H + L chains]) antibodies are usually
conjugated to horseradish peroxidase (HRPO), although other enzymes or conjugates (such as recombinant
protein G/HRPO) can be used. A monoclonal antibody to bovine IgG1–HRPO conjugate has also been found
to be suitable for use in the I-ELISA (Vigliocco et al., 1997). If a peroxidase conjugate is used, the
chromogen, usually ABTS, is diluted in a substrate buffer (composed of citric acid trisodium and citric acid;
pH 4). The substrate, hydrogen peroxide (H2O2), is added to this, and the plates are incubated for 15–
25 minutes at room temperature. The reaction may be stopped with 1 mM sodium azide, and the colour
change is read at 405–414 nm (for further details see chapter 2.4.3).
The antigen used in the I-ELISA is the HS in stock solution at 1 mg/ml in coating buffer, titrated in a
checkerboard titration, with different dilutions of antigen, conjugate and substrate, against a standard serum
or against serial dilutions of a panel of sera from B. ovis culture positive and Brucella-free sheep to
determine the most sensitive and specific dilution (usually 5–10 µg/ml).

Test procedure (example)
i)
Microtitre plates of good quality polystyrene are coated by the addition of 100 µl of a predetermined
antigen dilution in carbonate buffer, pH 9.6, to each well. Plates are incubated for 2 hours at 37°C.
Alternatively, the coating can be made overnight at 4°C with 100 µl/well of the predetermined antigen
dilution in PBS, pH 7.2. Plates are then washed four times to remove unbound antigen and dried by
tapping firmly upside down on an absorbent paper. The coated plates can be used immediately or
dried and stored at 4°C (the stability in these conditions is adequate for at least 1 month).
ii)
Sera: Dilute test and positive and negative control serum samples 1/200 by the addition of a minimum
of 10 µl of serum to 2 ml PBST. This serum dilution is usually the optimal when using both polyclonal
and monoclonal conjugates. However, dilutions of 1/50 are the optimal when using the protein G-
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
peroxidase conjugate (Marin et al., 1998) Add 100 µl/well volumes of samples in duplicate to the
microtitre plates. The plates are covered, incubated at 37°C for 1 hour, and washed three times with
the PBST washing buffer.
iii)
Conjugate: The titrated conjugate is diluted in PBST, added (100 µl) to the wells, and the plate is
covered and incubated for 1 hour at 37°C. After incubation, the plates are washed again three times
with PBST.
iv)
Substrate: The solution of ABTS in substrate buffer is added (100 µl/well) and the plates are incubated
for 15–60 minutes at room temperature with continuous shaking.
v)
Reading and interpreting the results: Absorbance is read automatically in a spectrophotometer at 405–
414 nm. Absorbance values may be expressed as percentages of the mean absorbance of the positive
control or, preferably, transformed into I-ELISA units calculated either manually or by using a computer
and a curve-fitting program from a standard curve constructed with the series of positive control dilution
results.
The cut-off threshold should be properly established using the appropriate validation techniques (see
Chapter 1.1.5 Principles and methods of validation of diagnostic assays for infectious diseases). The
International Standard for anti-Brucella ovis Serum or the corresponding secondary or national standards
should be used to check or calibrate the particular test method in question.
Comparative studies have shown that the I-ELISA has better sensitivity than either the AGID test or the CFT
(Marin et al., 1989; Ris et al., 1984; Spencer & Burgess, 1984; Worthington et al., 1984; 1985). Due to the
existence of some I-ELISA-negative and AGID-positive sera, the combination of the AGID test and I-ELISA
gives optimal sensitivity (Marin et al., 1989). However, the combination of CF test and I-ELISA or CF and
AGID tests does not improve the sensitivity of I-ELISA alone (Marin et al., 1989). Moreover, the CFT has
other important disadvantages such as complexity, obligatory serum inactivation, anticomplementary activity
of some sera, the difficulty of performing it with haemolysed sera, and prozone phenomena. Because of its
sensitivity, simplicity and easy interpretation, the AGID test is very practicable for routine diagnosis in
nonspecialised laboratories.
Little is known about the existence of false positive results in B. ovis serological tests as a consequence of
infections due to bacteria showing cross-reacting epitopes with B. ovis. The foot rot agent (Dichelobacter
nodosus) has been described as showing cross-reactions with B. ovis (Whitington et al., 1996), but the
extent and practical consequences of this cross-reactivity in B. ovis diagnostic tests is unknown4.
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICAIS
Vaccination of both rams and ewes is probably the most economical and practical means for medium-term control
of B. ovis in areas with a high incidence of infection. For long-term control, consideration should be given to the
effect of vaccination on serological testing, and B.-ovis-free accreditation programmes have to be implemented.
The live B. melitensis strain Rev.1 (see chapter 2.7.2) is probably the best available vaccine for the prophylaxis of
B. ovis infection (Blasco, 1990). A single standard dose (109 colony-forming units) of Rev.1 administered
subcutaneously (in a 1 ml volume) or conjunctivally (in a 25–30 µl volume), to 3–5 month-old animals confers
adequate immunity against B. ovis. Conjunctival vaccination has the advantage of minimising the intense and
long-lasting serological response evoked by subcutaneous vaccination, thereby improving the specificity of
serological tests (Blasco, 1990). When used in both young and adult males, the safety of the Rev.1 vaccine is
adequate and side-effects appear to be very rare (Marin et al., 1990; Muñoz et al., 2008). Therefore, in countries
with extensive management and high levels of incidence, it would be advisable to vaccinate both young and
healthy adult animals. In countries affected by B. ovis but free of B. melitensis, before using the B. melitensis
Rev.1 vaccine account should be taken of possible serological interferences and the conjunctival route should be
preferred to minimise this problem. The B. abortus RB51 live vaccine has not proven successful against B. ovis in
sheep (Jimenez et al., 1995) and no alternative vaccines are currently available.
REFERENCES
ALTON G.G., JONES L.M., ANGUS R.D. & VERGER J.M. (1988). Techniques for the Brucellosis Laboratory. INRA,
Paris, France.
4
Arcanobacterium pyogenes and Corynebacterium ovis, whose soluble extracts cross-react with B. ovis positive control
sera, have been recently isolated from several lymph nodes of rams giving strong positive responses in B. ovis AGID test
and I-ELISA (J.M. Blasco, unpublished results).
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
BLASCO J.M. (1990). Brucella ovis. In: Animal Brucellosis, Nielsen K. & Duncan J.R., eds. CRC Press, Boca
Raton, Florida, USA, 351–378.
BROWN G.M., RANGER C.R. & KELLEY D.J. (1971). Selective media for the isolation of Brucella ovis. Cornell Vet.,
61, 265–280.
BULGIN M.S. & ANDERSON B.C. (1983). Association of sexual experience with isolation of various bacteria in cases
of ovine epididymitis. J. Am. Vet. Med. Assoc., 182, 372–374.
BURGESS G.W. & MCDOWELL J.W. (1981). Escherichia coli epididymitis and seminal vesiculitis in a ram. Aust. Vet.
J., 57, 479–480.
BURGESS G.W. & NORRIS M.J. (1982). Evaluation of the cold complement fixation test for diagnosis of ovine
brucellosis. Aust. Vet. J., 59, 23–25.
CORBEL M.J., GILL K.P.W. & THOMAS E.L. (1978). Methods for the identification of Brucella. Ministry of Agriculture,
Fisheries and Food, UK, ADAS, RCV 22.
DE LONG W.J., WALDHALM D.G. & HALL R.F. (1979). Bacterial isolates associated with epididymitis in rams from
Idaho and eastern Oregon flocks. Am. J. Vet. Res., 40, 101–102.
DIAZ R. & BOSSERAY N. (1973). Identification d’un composé antigénique spécifique de la phase rugueuse (R) des
Brucella. Ann. Rech. Vet., 4, 283–292.
EKDAHL M.O., MONEY D.F. & MARTIN C.A. (1968). Some aspects of epididymitis of rams in New Zealand. N. Z. Vet.
J., 16, 81–82.
JIMENEZ DE BAGUES M.P., BARBERAN M., MARIN C.M. & BLASCO J.M. (1995). The Brucella abortus RB51 vaccine
does not confer protection against Brucella ovis in rams. Vaccine, 13, 301–304.
LIVINGSTONE C.W. & HARDY W.T. (1964). Isolation of Actinobacillus seminis from ovine epididymitis. Am. J. Vet.
Res., 25, 660–663.
MARIN C.M., ALABART J.L. & BLASCO J.M. (1996). Effect of antibiotics contained in two Brucella selective media on
growth of B. abortus, B. melitensis and B. ovis. J. Clin. Microbiol., 34, 426–428.
MARIN C.M., ALONSO-URMENETA B., MORIYON I., PEREZ S. & BLASCO J.M. (1998). Comparison of polyclonal,
monoclonal and protein G peroxidase conjugates in an enzyme-linked immunosorbent assay for the diagnosis of
Brucella ovis in sheep. Vet. Rec., 143, 390–394.
MARIN C.M., BARBERAN M., JIMENEZ DE BAGUES M.P. & BLASCO J.M. (1990). Comparison of subcutaneous and
conjunctival routes of Rev. 1 vaccination for the prophylaxis of Brucella ovis infection in rams. Res. Vet. Sci., 48,
209–215.
MARIN C.M., JIMENEZ DE BAGUES M.P., BLASCO J.M., GAMAZO C., MORIYON I. & DIAZ R. (1989). Comparison of three
serological tests for Brucella ovis infection of rams using different antigenic extracts. Vet. Rec., 125, 504–508.
MUÑOZ P., DE MIGUEL M.J., GRILLÓ M.J., MARÍN C.M., BARBERÁN M. & BLASCO J.M. (2008). Immunopathological
responses and kinetics of B. melitensis Rev 1 infection after subcutaneous or conjunctival vaccination in rams.
Vaccine, 26, 2562–2569.
MYERS D.M., JONES L.M. & VARELA-DIAZ V. (1972). Studies of antigens for complement fixation and gel diffusion
tests in the diagnosis of infections caused by Brucella ovis and other Brucella. Appl. Microbiol., 23, 894–902.
RIDLER A. (2001) Brucella ovis infection in deer. Surveillance, 28 (3), 6–8.
RIEZU-BOJ J.I., MORIYON I., BLASCO J.M., MARIN C.M. & DIAZ R. (1986). Comparison of lipopolysaccharide and outer
membrane protein-lipopolysaccharide extracts in an enzyme-linked immunosorbent assay for the diagnosis of
Brucella ovis infection. J. Clin. Microbiol., 23, 938–942.
RIS D.R., HAMEL K.L. & LONG D.L. (1984). Comparison of an enzyme-linked immunospecific assay (ELISA) with the
cold complement fixation test for the serodiagnosis of Brucella ovis infection. N.Z. Vet. J., 32, 18–20.
OIE Terrestrial Manual 2012
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Chapter 2.7.9. — Ovine epididymitis (Brucella ovis)
RODOLAKIS A. & BERNARD K. (1977). Isolement de Chlamydia des organes génitaux de béliers atteints
d’épididymite. Bull. Acad. Vet. Fr., 50, 65–70.
SANTOS J.M., VERSTREATE D.R., PERERA V.Y. & WINTER A.J. (1984). Outer membrane proteins from rough strains of
four Brucella species. Infect. Immun., 46, 188–194.
SEARSON J.E. (1982). Sensitivity and specificity of two microtitre complement fixation tests for the diagnosis of
Brucella ovis infection in rams. Aust. Vet. J., 58, 5–7.
SPENCER T.L. & BURGESS G.W. (1984). Enzyme-linked immunosorbent assay for Brucella ovis specific antibody in
ram sera. Res. Vet. Sci., 36, 194–198.
VIGLIOCCO A.M., SILVA PAULO P.S., MESTRE J., BRIONES G.C., DRAGHI G., TOSSI M. & NIELSEN K. (1997).
Development and validation of an indirect enzyme immunoassay for detection of ovine antibody to Brucella ovis.
Vet. Microbiol., 54, 357–368.
WEBB R.F. (1983). Clinical findings and pathological changes in Histophilus ovis infections of sheep. Res. Vet.
Sci., 35, 30–34.
WEBB R.F., QUINN C.A., COCKRAM F.A. & HUSBAND A.J. (1980). Evaluation of procedures for the diagnosis of
Brucella ovis infection in rams. Aust. Vet. J., 56, 172–175.
WHITINGTON R.J., SAUNDERS V.F. & EGERTON J.R. (1996). Antigenic cross-reactions between the causative agent of
ovine footrot, Dichelobacter nodosus, and other bacteria. Small Rumin. Res., 22, 55–67.
WILLIAMSON P. & NAIRN M.E. (1980). Lesions caused by Corynebacterium pseudotuberculosis in the scrotum of
rams. Aust. Vet. J., 56, 496–498.
WORTHINGTON R.W., STEVENSON B.J. & DE LISLE G.W. (1985). Serology and semen culture for the diagnosis of
Brucella ovis infection in chronically infected rams. N.Z. Vet. J., 33, 84–86.
WORTHINGTON R.W., WEDDELL W. & PENROSE M.E. (1984). A comparison of three serological tests for the diagnosis
of B. ovis infection in rams. N.Z. Vet. J., 32, 58–60.
*
* *
NB: There are OIE Reference Laboratories for Ovine epididymitis (Brucella ovis)
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for ovine epididymitis (Brucella ovis)
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2008
CHAPTER 2.7.10.
OVINE PULMONARY ADENOCARCINOMA
(adenomatosis)
SUMMARY
Ovine pulmonary adenocarcinoma (OPA), also known as ovine pulmonary adenomatosis and
jaagsiekte, is a contagious tumour of sheep and, rarely, of goats. It is a progressive respiratory
disease, principally affecting adult animals. The disease occurs in many regions of the world. A
beta-retrovirus (jaagsiekte sheep retrovirus: JSRV), distinct from the non-oncogenic ovine
lentiviruses, has been shown to cause the disease.
Identification of the agent: JSRV cannot yet be propagated in vitro, therefore routine diagnostic
methods, such as virus isolation, are not available for diagnosis. Diagnosis relies, at present, on
clinical history and examination, as well as on the findings at necropsy and by histopathology and
immunohistochemistry. Viral DNA or RNA can be detected in tumour, draining lymph nodes, and
peripheral blood mononuclear cells by polymerase chain reaction. Lambs become persistently
infected by JSRV at an early age, and, in an OPA-affected flock, most sheep are infected.
Serological tests: Antibodies to the retrovirus have not been detected in infected sheep and,
therefore, serological tests are not available for diagnosis.
Requirements for vaccines and diagnostic biologicals: There are no vaccines or diagnostic
biologicals available.
A. INTRODUCTION
Ovine pulmonary adenocarcinoma (OPA), also known as ovine pulmonary adenomatosis, jaagsiekte (Afrikaans =
driving sickness) and ovine pulmonary carcinoma (OPC), is a contagious lung tumour of sheep and, more rarely,
of goats. It is the most common pulmonary tumour of sheep and occurs in many countries around the world. It is
absent from Australia and New Zealand and has been eradicated from Iceland.
A number of different viruses have been linked aetiologically to OPA, including a herpesvirus and lentiviruses that
have been propagated from tumour tissue. However, the former does not have an aetiological role in OPA and
the latter exhibit characteristics of non-oncogenic lentiviruses. It has been demonstrated clearly that OPA is
caused by a beta-retrovirus that cannot yet be cultured in vitro, but the virus has been cloned and sequenced.
The term jaagsiekte sheep retrovirus (JSRV) is used in referring to this virus.
B. DIAGNOSTIC TECHNIQUES
At present, diagnosis of OPA relies on clinical and pathological investigations, although polymerase chain
reaction (PCR) offers hope for ante-mortem diagnosis of OPA as a flock test. In flocks in which the disease is
suspected, its presence must be, at least once, confirmed by histopathological examination of affected lung
tissue. For such an examination, it is imperative to take specimens from several affected sites and, if possible,
from more than one animal. This is because secondary bacterial pneumonia, which might be the immediate cause
of death, often masks the lesions (both macroscopic and microscopic) of the primary disease. In the absence of
specific serological tests that can be used for the diagnosis of OPA in live animals, disease control relies on
regular flock inspections and prompt culling of suspected cases and, in the case of ewes, their offspring.
1.
Identification of the agent
Although ovine herpesvirus 1 (OvHV-1) had been isolated exclusively from OPA tumours, epidemiological studies
and experimental infections provide no evidence for a role in the aetiology of OPA. Ovine herpesvirus 2 (OvHV-2)
is the sheep-associated malignant catarrhal fever herpesvirus and has never been linked to OPA.
OIE Terrestrial Manual 2012
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Chapter 2.7.10. — Ovine pulmonary adenomatosis (adenocarcinoma)
The association of retroviruses with OPA has been recognised for several years. Ovine lentiviruses have been
isolated on a number of occasions, but these viruses have no aetiological role in OPA.
For many years, the inability to culture JSRV and the lack of antibodies to the virus in affected sheep impeded the
confirmation of this virus as the aetiological agent. However, molecular biological techniques provided a key
advance, namely, the cloning and sequencing of the 7.5 kb JSRV genome following purification of virions from
lung washes of naturally affected sheep (York et al., 1992). JSRV has been designated as a beta-retrovirus
because of its genetic organisation and its structural proteins. Although cloned JSRV genes, used as hybridisation
probes, have revealed a range of homologous endogenous sequences in the genome of both healthy and OPAaffected sheep (Bai et al., 1996; Hecht et al., 1994; York et al., 1992), JSRV is clearly exogenous and associated
exclusively with OPA (Palmarini et al., 1996). JSRV is detected constantly in the lung fluid, tumour, peripheral
blood mononuclear cells, and lymphoid tissues of sheep affected by OPA or unaffected in-contact flockmates, and
never in sheep from unaffected flocks with no history of the tumour. Full-length proviral clones of JSRV have been
obtained from OPA tumour DNA and cells. JSRV virus particles, prepared from these clones by transient
transfection of a cell line, were used for intratracheal inoculation of neonatal lambs. OPA tumour was induced in
the lambs, thus demonstrating that JSRV is the causal agent of OPA (DeMartini et al., 2001; Palmarini et al.,
1999).
The sheep genome contains many copies of endogenous viral sequences that are highly related to JSRV.
Although they are not involved in the aetiology of OPA, their expression in the fetus may, by induction of
tolerance, account for the apparent lack of immune response of mature animals to exogenous JSRV (Palmarini et
al., 2004).
a)
Nucleic acid recognition methods
Sequencing of JSRV and endogenous sequences in the sheep genome has led to the development of PCRs
that specifically detect JSRV (Bai et al., 1996; Palmarini et al., 1996). Using this sensitive procedure, JSRV
has been detected in peripheral blood mononuclear cells of unaffected in-contact sheep from flocks with
OPA, as well as experimentally infected lambs (De las Heras et al., 2005; Gonzalez et al., 2001; Holland et
al., 1999) and the bronchoalveolar lavage samples from unaffected in-contact sheep (Voigt et al., 2007).
Longitudinal studies in OPA-affected flocks have shown that lambs become infected at a very early age. A
high proportion of animals in these flocks are infected, yet only a minority develop OPA (Caporale et al.,
2005; Salvatori, 2005). JSRV has been found in colostrum and milk obtained from sheep in OPA-affected
flocks and JSRV can be detected within a few months in the blood of lambs fed artificially with colostrum and
milk (De las Heras et al., unpublished observations).

Control and treatment
b)
Animal inoculation
OPA cannot be transmitted to any laboratory animal and can be transmitted to sheep only with material that
contains JSRV, such as tumour homogenates, concentrated cell-free lung fluid from natural cases of OPA
and virus produced from molecular clones. Following the experimental inoculation of adult sheep, clinical
disease develops only after several months or years. In contrast, JSRV infection can be induced in 100% of
lambs aged 1–6 months at the time of inoculation and a high proportion of these animals develop clinical
signs (62–90%) and lesions (87–100%) of OPA (Salvatori et al., 2004).
At this time there is no practical animal inoculation method for the diagnosis of OPA.
c)
Virus isolation
There are no permissive cell culture systems for propagation of JSRV. Some cell cultures prepared from the
tumours occurring in young lambs can support virus replication for a short period (Jassim, 1988; Sharp et al.,
1985).
d)
Clinical signs and pathology
There is no reliable laboratory method for the ante-mortem diagnosis of OPA in individual animals at this
time, therefore flock history, clinical signs and post-mortem lesions are the primary method for the diagnosis
of the disease. As OPA has a long incubation period, clinical disease is encountered most commonly in
sheep over 2 years of age, with a peak occurrence at the age of 3–4 years. In exceptional cases, the
disease occurs in animals as young as 2–3 months of age. The cardinal signs are those of a progressive
respiratory embarrassment, particularly after exercise; the severity of the signs reflects the extent of tumour
development in the lungs. Accumulation of fluid within the respiratory tract is a prominent feature of OPA,
giving rise to moist râles that are readily detected by auscultation. Raising the hindquarters and lowering the
head of affected sheep may cause frothy mucoid fluid to run from the nostrils. Coughing and inappetance
are not common but, once clinical signs are evident, weight loss is progressive and the disease is terminal
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Chapter 2.7.10. — Ovine pulmonary adenocarcinoma (adenomatosis)
within weeks or months. Death is often precipitated by a superimposed bacterial pneumonia, particularly that
due to Mannheimia (formerly Pasteurella) haemolytica. In clinically affected animals, a peripheral
lymphopenia characterised by a reduction in CD4+ T lymphocytes and a corresponding neutrophilia may
assist clinical diagnosis, but the changes are not pathognomonic and are not detected during early
experimental infection (Summers et al., 2002).
In some countries, another form of OPA (atypical OPA) occurs, which generally presents as an incidental
finding at necropsy or the abattoir (De las Heras et al., 2003).
e)
Necropsy
OPA lesions are in most cases confined to the lungs, although intra- and extrathoracic metastasis to lymph
nodes and other tissues can occur. In typical cases, affected lungs are considerably enlarged and heavier
than normal due to extensive nodular and coalescing firm grey lesions affecting much of the pulmonary
tissue. Usually lesions are present in both lungs, although the extent on either side does vary. Tumours are
solid, grey or light purple with a shiny translucent sheen and often separated from the adjacent normal lung
by a narrow emphysematous zone. The presence of frothy white fluid in the respiratory passages is a
prominent feature and is obvious even in lesions as small as a few millimetres. In advanced cases, this fluid
flows out of the trachea when it is cut or pendant. Samples should be taken at necropsy for histopathology,
immunohistochemistry or PCR for JSRV.
Pleurisy may be evident over the surface of the tumour and often abscesses are present in the
adenomatous tissue.
In atypical OPA, tumours comprise solitary or aggregated hard white nodules that have a dry cut surface and
show clear demarcation from surrounding tissues. The presence of excess fluid is not a prominent feature.
Adult sheep, which on post-mortem examination appear to have died from acute pasteurellosis, should have
their lungs examined carefully, as lesions of OPA may be masked by coexisting bronchopneumonia,
verminous pneumonia, chronic progressive pneumonia (maedi-visna) or combinations of these. Samples
should be taken at necropsy for histopathology.
f)
Histopathology
Histologically, the lesions are characterised by proliferation of mainly type II pneumocytes, a secretory
epithelial cell in the pulmonary alveoli. Nonciliated (Clara) and epithelial cells of the terminal bronchioli may
be involved. The cuboidal or columnar tumour cells replace the normal thin alveolar cells and sometimes
form papilliform growths that project into the alveoli. Intrabronchiolar proliferation may be present. In
advanced cases, extensive fibrosis may develop and, occasionally, nodules of loose connective tissue in a
mucopolysaccharide substance may be present.
A prominent feature is the accumulation of large numbers of alveolar macrophages in the alveoli adjacent to
the neoplastic lesions (Summers et al., 2005).
Where maedi-visna is concurrent, perivascular, peribronchiolar and interstitial lymphoid infilrates may be
prominent.
The histological appearance of atypical OPA is essentially the same as classical OPA, but with an
exaggerated inflammatory response (mostly lymphocytes and plasma cells) and fibrosis (De las Heras et al.,
2003).
For more detailed accounts of the clinical, post-mortem and histopathological aspects of OPA, the reader is
referred elsewhere (De las Heras et al., 2003; Sharp & DeMartini, 2003).
There appears to be a synergistic interaction between OPA and maedi-visna. Lateral transmission of maedivisna virus appears to be enhanced in sheep affected by OPA (Dawson et al., 1985; Gonzalez et al., 1993).
2.
Serological tests
At present, there are no laboratory tests to support a clinical diagnosis of OPA in the live animal. JSRV has been
associated exclusively with both typical and atypical forms of OPA, but antibodies to this virus have not been
detected in the sera of affected sheep, even with highly sensitive assays such as immunoblotting or enzymelinked immunosorbent assay (Ortin et al., 1997; Summers et al., 2002).
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Chapter 2.7.10. — Ovine pulmonary adenomatosis (adenocarcinoma)
C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
There are no vaccines or diagnostic biologicals available at the present time.
REFERENCES
BAI J., ZHU R.-Y., STEDMAN K., COUSENS C., CARLSON J.O., SHARP J.M. & DEMARTINI J.C. (1996). Unique long
terminal repeat U3 sequences distinguish exogenous jaagsiekte sheep retroviruses associated with ovine
pulmonary carcinoma from endogenous loci in the sheep genome. J. Virol., 70, 3159–3168.
CAPORALE M., CENTORAME P., GIOVANNINI A., SACCHINI F., DI VENTURA M., DE LAS HERAS M. & PALMARINI M. (2005).
Infection of lung epithelial cells and induction of pulmonary adenocarcinoma is not the most common outcome of
naturally occurring JSRV infection during the commercial lifespan of sheep. Virology, 338, 144–153.
DAWSON M., VENABLES C. & JENKINS C.E. (1985). Experimental infection of a natural case of sheep pulmonary
adenomatosis with maedi-visna virus. Vet. Rec. 116, 588–589.
DE LAS HERAS M., GONZALEZ L.G. & SHARP J.M. (2003). Pathology of ovine pulmonary adenocarcinoma. Curr. Top.
Microbiol. Immunol., 275, 25–54
DE LAS HERAS M., ORTÍN A., SALVATORI D., PÉREZ DE VILLAREAL M., COUSENS C., FERRER L.M., GARCÍA DE JALÓN J.A.,
GONZALEZ L. & SHARP J.M. (2005). A PCR technique for the detection of Jaagsiekte retrovirus in the blood suitable
for the screening of virus infection in sheep flocks. Res. Vet. Sci., 79, 259–264.
DEMARTINI J.C., BISHOP J.V., ALLEN T.E., JASSIM F.A., SHARP J.M., DE LAS HERAS M., VOELKER D.R. & CARLSON J.O.
(2001). Jaagsiekte sheep retrovirus proviral clone JSRVJS7, derived from the JS7 lung tumor cell line, induces
ovine pulmonary carcinoma and is integrated into the surfactant protein A gene. J. Virol., 75, 4239–4246
GONZALEZ L., GARCIA-GOTI M., COUSENS C., DEWAR P., CORTABARRIA N., EXTRAMIANA B., ORTIN A., DE LAS HERAS M.
& SHARP J.M. (2001). Jaagsiekte sheep retrovirus can be detected in the peripheral blood during the preclinical
period of sheep pulmonary adenomatosis. J. Gen. Virol., 82, 1355–1358.
GONZALEZ L., JUSTE R.A., CUERVO L.A., IDIGORAS I. & SAEZ DE OCARIZ C. (1993). Pathological and epidemiological
aspects of the coexistence of maedi-visna and sheep pulmonary adenomatosis. Res. Vet. Sci., 54, 140–146.
HECHT S.J., CARLSON J.O. & DE MARTINI J.C. (1994). Analysis of a type D retroviral capsid gene expressed in ovine
pulmonary carcinoma and present in both affected and unaffected sheep genomes. Virology, 202, 480–484.
HOLLAND M.J., PALMARINI M., GARCIA-GOTI M., GONZALEZ L., DE LAS HERAS M. & SHARP J.M. (1999). Jaagsiekte
retrovirus establishes a pantropic infection of lymphoid cells of sheep with naturally and experimentally acquired
pulmonary adenomatosis. J. Virol., 73, 4004–4008.
JASSIM F.A. (1988). Identification and characterisation of transformed cells in jaagsiekte, a contagious lung tumour
of sheep. PhD thesis. University of Edinburgh, UK.
ORTIN A., MINGUIJON E., DEWAR P., GARCIA M., FERRER L.M., PALMARINI M., GONZALEZ L., SHARP J.M. & DE LAS HERAS
M. (1997). Lack of a specific immune response against a recombinant capsid protein of Jaagsiekte sheep
retrovirus in sheep and goats naturally affected by enzootic nasal tumour or sheep pulmonary adenomatosis. Vet.
Immunol. Immunopathol., 61, 239–237.
PALMARINI M., COUSENS C., DALZIEL R.G., BAI J., STEDMAN K, DEMARTINI J.C. & SHARP J.M. (1996). The exogenous
form of Jaagsiekte retrovirus (JSRV) is specifically associated with a contagious lung cancer of sheep. J. Virol.,
70, 1618–1623.
PALMARININ M., MURA M. & SPENCER T. (2004). Endogenous betaretroviruses of sheep: teaching new lessons in
retroviral interference and adaptation. J. Gen. Virol., 85, 1–13.
PALMARINI M., SHARP J.M., DE LAS HERAS M. & FAN H.Y. (1999). Jaagsiekte sheep retrovirus is necessary and
sufficient to induce a contagious lung cancer in sheep. J. Virol., 73, 6964–6972.
SALVATORI D. (2005). Studies on the pathogenesis and epidemiology of ovine pulmonary adenomatosis (OPA).
PhD thesis, University of Edinburgh, Scotland, UK.
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SALVATORI D., COUSENS C., DEWAR P., ORTIN A., GONZALEZ L., DE LAS HERAS M., DALZIEL R.G. & SHARP J.M. (2004).
Effect of age at inoculation on the development of ovine pulmonary adenocarcinoma. J. Gen. Virol., 85, 3319–
3324.
SHARP J.M. & DEMARTINI J.C. (2003). Natural history of JSRV in sheep. Curr. Top. Microbiol. Immunol., 275, 55–
79.
SHARP J.M., HERRING A.J., ANGUS K.W., SCOTT F.M.M. & JASSIM F.A. (1985). Isolation and in vitro propagation of a
retrovirus from sheep pulmonary adenomatosis. In: Slow Virus Diseases in Sheep, Goats and Cattle. Sharp J.M.
& Hoff-Jorgensen R., eds. CEC Report EUR 8076 EN, Luxembourg, 345–348.
SUMMERS C., NEILL W., DEWAR P., GONZALEZ L., VAN DER MOLEN R., NORVAL M. & SHARP J.M. (2002). Systemic
immune responses following infection with jaagsiekte sheep retrovirus and in the terminal stages of ovine
pulmonary adenocarcinoma. J. Gen. Virol., 83, 1753–1757.
SUMMERS C., NORVAL M., DE LAS HERAS M., GONZALEZ L., SHARP J.M. & WOODS G.M. (2005). An influx of
macrophages is the predominant local immune response in ovine pulmonary adenocarcinoma. Vet. Immunol.
Immunopathol., 106, 285–294.
VOIGT K., BRÜGMANN M., HUBER K., DEWAR P., COUSENS C., HALL M., SHARP J.M. & GANTER M. (2007). PCR
examination of bronchoalveolar lavage samples is a useful tool in pre-clinical diagnosis of ovine pulmonary
adenocarcinoma (Jaagsiekte). Res. Vet. Sci., 83 (3), 419–427.
YORK D.F., VIGNE R., VERWOERD D.W. & QUERAT G. (1992). Nucleotide sequence of the jaagsiekte retrovirus, an
exogenous and endogenous type D and B retrovirus of sheep and goats. J. Virol., 66, 4930–4939.
*
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2012
CHAPTER 2.7.11.
PESTE DES PETITS RUMINANTS
SUMMARY
Peste des petits ruminants (PPR), is an acute contagious disease caused by a Morbillivirus in the
family Paramyxoviridae. It affects mainly sheep and goats and occasionally wild small ruminants.
Based on the fact that PPR has been reported on a few occasions in camels, cattle and buffaloes,
those animal species are considered to be susceptible although their potential role in the circulation
of PPR virus (PPRV) has not been formally established. PPR occurs in Africa except Southern
Africa, in the Arabian Peninsula, throughout most of the Near East and Middle East, and in Central
and South-East Asia.
The clinical disease resembles rinderpest in cattle. It is usually acute and characterised by pyrexia,
serous ocular and nasal discharges, erosive lesions on different mucous membranes particularly in
the mouth, diarrhoea and pneumonia. At necropsy, erosions may be noted in the gastrointestinal
and urogenital tracts. The lungs may show interstitial bronchopneumonia and often secondary
bacterial pneumonia. PPR can also occur in subclinical form.
The disease must be differentiated from rinderpest, bluetongue, foot and mouth disease and other
exanthemous conditions.
Identification of the agent: The collection of specimens at the correct time is important to achieve
diagnosis by virus isolation and they should be obtained in the acute phase of the disease when
clinical signs are still apparent. The specimens from live animals can be swabs of conjunctival
discharges, nasal secretions, buccal and rectal mucosae, and anticoagulant-treated blood.
Rapid diagnosis is done by immunocapture enzyme-linked immunosorbent assay (ELISA), counter
immunoelectrophoresis and agar gel immunodiffusion. Polymerase chain reaction may also be
used.
Serological tests: The serological tests that are routinely used are the virus neutralisation and the
competitive ELISA.
Requirements for vaccines: In the past, control of PPR was ensured through vaccination with the
rinderpest tissue culture vaccine because of the existence of a strong antigenic relationship
between PPR and rinderpest viruses. The use of this heterologous vaccine has been abandoned in
favour of the live attenuated PPR virus vaccine, which is now widely commercially available.
A. INTRODUCTION
Peste des petits ruminants (PPR) is an acute viral disease of small ruminants characterised by fever, oculonasal
discharges, stomatitis, diarrhoea and pneumonia with foul offensive breath. Because of the respiratory signs, PPR
can be confused with contagious caprine pleuropneumonia (CCPP) or pasteurellosis. In many cases,
pasteurellosis is a secondary infection of PPR, a consequence of the immunosuppression that is induced by the
PPR virus (PPRV). PPRV is transmitted mainly by aerosols between animals living in close contact (Lefevre &
Diallo, 1990). Infected animals present clinical signs similar to those historically seen with rinderpest in cattle,
although the two diseases are caused by distinct virus species.
On the basis of its similarities to rinderpest, canine distemper and measles viruses, PPRV has been classified
within the genus Morbillivirus in the family Paramyxoviridae (Gibbs et al., 1979). Virus members of this group
have six structural proteins: the nucleocapsid protein (Np), which encapsulates the virus genomic RNA, the
phosphoprotein (P), which associates with the polymerase (L for large protein) protein, the matrix (M) protein, the
fusion (F) protein and the haemagglutinin (H) protein. The matrix protein, intimately associated with the internal
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Chapter 2.7.11. — Peste des petits ruminants
face of the viral envelope, makes a link between the nucleocapsid and the virus external glycoproteins: H and F,
which are responsible, respectively, for the attachment and the penetration of the virus into the cell to be infected.
PPR was first described in Côte d’Ivoire (Gargadennec & Lalanne, 1942), but it occurs in most African countries
from North Africa to Tanzania (Kwiatek et al., 2011; Lefevre & Diallo, 1990; Swai, et al., 2009), and in nearly all
Middle Eastern countries to Turkey (Furley et al., 1987; Lefevre et al., 1991; Ozkul et al., 2002; Perl et al., 1994;
Taylor et al., 1990). PPR is also widespread in countries from a central Asia to South and South-East Asia
(Banyard et al., 2010; Shaila et al., 1989; Wang et al., 2009).
The natural disease affects mainly goats and sheep. It is generally admitted that cattle can only be infected
subclinically. However, in poor conditions it might be possible for cattle to develop lesions following PPRV
infection, clinical signs of which would be ascribed to rinderpest. Indeed, in the 1950s, disease and death were
recorded in calves experimentally infected with PPRV-infected tissue (Mornet et al., 1956). In addition, the virus
that was involved in what was thought to be a natural rinderpest outbreak in cattle, sheep and goats in 1971 in
Sudan was later shown to be PPRV (Kwiatek et al., 2011). Moreover, PPRV was isolated from an outbreak of
rinderpest-like disease in buffaloes in India in 1995 (Govindarajan et al., 1997). Antibodies to PPRV as well as
PPRV antigen and nucleic acid were detected in some samples from an epizootic disease that affected onehumped camels in Ethiopia and Sudan (Abraham et al., 2005; Ismail et al., 1992; Khalafalla et al. 2010; Kwiatek
et al., 2011; Roger et al., 2000; 2001). Cases of clinical disease have been reported in wildlife resulting in deaths
of wild small ruminants (Abubakar et al., 2011; Bao et al., 2011; Elzein et al., 2004; Furley et al., 1987; Kinne et
al., 2010). The American white-tailed deer (Odocoileus virginianus) can be infected experimentally with PPRV
(Hamdy & Dardiri, 1976). Dual infections can occur with other viruses such as pestivirus or goatpox virus.
The incubation period is typically 4–6 days, but may range between 3 and 10 days. The clinical disease is acute,
with a pyrexia up to 41°C that can last for 3–5 days; the animals become depressed, anorexic and develop a dry
muzzle. Serous oculonasal discharges become progressively mucopurulent and, if death does not ensue, persist
for around 14 days. Within 4 days of the onset of fever, the gums become hyperaemic, and erosive lesions
develop in the oral cavity with excessive salivation. These lesions may become necrotic. A watery blood-stained
diarrhoea is common in the later stage. Pneumonia, coughing, pleural rales and abdominal breathing also occur.
The morbidity rate can be up to 100% with very high case fatality in severe cases. However, morbidity and
mortality may be much lower in milder outbreaks, and the disease may be overlooked. A tentative diagnosis of
PPR can be made on clinical signs, but this diagnosis is considered provisional until laboratory confirmation is
made for differential diagnosis with other diseases with similar signs.
At necropsy, the lesions are very similar to those observed in cattle affected with rinderpest, except that
prominent crusty scabs along the outer lips and severe interstitial pneumonia frequently occur with PPR. Erosive
lesions may extend from the mouth to the reticulo–rumen junction. Characteristic linear red areas of congestion or
haemorrhage may occur along the longitudinal mucosal folds of the large intestine and rectum (zebra stripes), but
they are not a consistent finding. Erosive or haemorrhagic enteritis is usually present and the ileocaecal junction
is commonly involved. Peyer’s patches may be necrotic. Lymph nodes are enlarged, and the spleen and liver may
show necrotic lesions.
There are no known health risks to humans working with PPRV as no report of human infection with the virus
exists.
B. DIAGNOSTIC TECHNIQUES
1.
Collection of samples
Samples for virus isolation must be kept chilled in transit to the laboratory. In live animals, swabs are made of the
conjunctival discharges and from the nasal and buccal mucosae. During the very early stage of the disease,
whole blood is also collected in anticoagulant for virus isolation, polymerase chain reaction (PCR) and
haematology. At necropsy, samples from two to three animals should be collected aseptically from lymph nodes,
especially the mesenteric and bronchial nodes, lungs, spleen and intestinal mucosae, chilled on ice and
transported under refrigeration. Samples of organs collected for histopathology are placed in 10% neutral buffered
formalin. It is good practice to collect blood for serological diagnosis at all stages, but particularly later in the
outbreak.
2.
Identification of the agent
a)
Agar gel immunodiffusion
Agar gel immunodiffusion (AGID) is a very simple and inexpensive test that can be performed in any
laboratory and even in the field. Standard PPR viral antigen is prepared from infected mesenteric or
bronchial lymph nodes, spleen or lung material and ground up as 1/3 suspensions (w/v) in buffered saline
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Chapter 2.7.11. — Peste des petits ruminants
(Durojaiye et al., 1983). These are centrifuged at 500 g for 10–20 minutes, and the supernatant fluids are
stored in aliquots at –20°C. The cotton material from the cotton bud used to collect eye or nasal swabs is
removed using a scalpel and inserted into a 1 ml syringe. With 0.2 ml of phosphate buffered saline (PBS),
the sample is extracted by repeatedly expelling and filling the 0.2 ml of PBS into an Eppendorf tube using
the syringe plunger. The resulting eye/nasal swab extracted sample, like the tissue ground material prepared
above, may be stored at –20°C until used. They may be retained for 1–3 years. Negative control antigen is
prepared similarly from normal tissues. Standard antiserum is made by hyperimmunising sheep with 1 ml of
PPRV with a titre of 104 TCID50 (50% tissue culture infective dose) per ml given at weekly intervals for
4 weeks. The animals are bled 5–7 days after the last injection (Durojaiye, 1982).
i)
Dispense 1% agar in normal saline, containing thiomersal (0.4 g/litre) or sodium azide (1.25 g/litre) as a
bacteriostatic agent, into Petri dishes (6 ml/5 cm dish).
ii)
Six wells are punched in the agar following a hexagonal pattern with a central well. The wells are 5 mm
in diameter and 5 mm apart.
iii)
The central well is filled with positive antiserum, three peripheral wells with positive antigen, and one
well with negative antigen. The two remaining peripheral wells are filled with test antigen, such that the
test and negative control antigens alternate with the positive control antigens.
iv)
Usually, 1–3 precipitin lines will develop between the serum and antigens within 18–24 hours at room
temperature (Durojaiye et al., 1983). These are intensified by washing the agar with 5% glacial acetic
acid for 5 minutes (this procedure should be carried out with all apparently negative tests before
recording a negative result). Positive reactions show lines of identity with the positive control antigen.
Results are obtained in one day, but the test is not sensitive enough to detect mild forms of PPR due to the
low quantity of viral antigen that is excreted.
b)
Counter immunoelectrophoresis
Counter immunoelectrophoresis (CIEP) is the most rapid test for viral antigen detection (Majiyagbe et al.,
1984). It is carried out on a horizontal surface using a suitable electrophoresis bath, which consists of two
compartments connected through a bridge. The apparatus is connected to a high-voltage source. Agar or
agarose (1–2%, [w/v]) dissolved in 0.025 M barbitone acetate buffer is dispensed on to microscope slides in
3-ml volumes. From six to nine pairs of wells are punched in the solidified agar. The reagents are the same
as those used for the AGID test. The electrophoresis bath is filled with 0.1 M barbitone acetate buffer. The
pairs of wells in the agar are filled with the reactants: sera in the anodal wells and antigen in the cathodal
wells. The slide is placed on the connecting bridge and the ends are connected to the buffer in the troughs
by wetted porous paper. The apparatus is covered, and a current of 10–12 milliamps per slide is applied for
30–60 minutes. The current is switched off and the slides are viewed by intense light: the presence of 1–
3 precipitation lines between pairs of wells is a positive reaction. There should be no reactions between
wells containing the negative controls.
c)
Immunocapture enzyme-linked immunosorbent assay
Advice on the use and applicability of ELISA methods is available from the OIE Reference Laboratories for
PPR. Both the methods described are available as commercial kits.
The immunocapture enzyme-linked immunosorbent assay (ELISA) (Libeau et al., 1994) using two
monoclonal antibodies (MAb) raised to the N protein, allows a rapid identification of PPRV. The instructions
provided by kit supplier should be followed, but the following shows a typical procedure for the test.
i)
Microtitre ELISA plates (such as Nunc Maxisorp) are coated with 100 µl of a capture MAb solution
(diluted according to the instructions of the kit supplier). Coating may be overnight at 4°C or for 1 hour
at 37°C.
ii)
After washing, 50 µl of the sample suspension is added to two wells, and control wells are filled with
buffer.
iii)
Immediately, add 25 µl of a detection biotinylated MAb for PPR and 25 µl of streptavidin/peroxidase to
two wells.
iv)
The plates are incubated at 37°C for 1 hour with constant agitation.
v)
After three vigorous washes, 100 µl of ortho-phenylenediamine (OPD) in hydrogen peroxide is added,
and the plates are incubated for 10 minutes at room temperature.
vi)
The reaction is stopped by the addition of 100 µl of 1 N sulphuric acid, and the absorbance is
measured at 492 nm on a spectrophotometer/ELISA reader.
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Chapter 2.7.11. — Peste des petits ruminants
The cut-off above which samples are considered to be positive is calculated from the blank as three times
the mean absorbance values.
The test is very specific and sensitive (it can detect 100.6 TCID50/well of PPRV). The results are obtained in
2 hours.
A sandwich form of the immunocapture ELISA is widely used in India (Singh et al., 2004a): the sample is
first allowed to react with the detection MAb and the immunocomplex is then captured by the MAb or
polyclonal antibody adsorbed on to the ELISA plate. The assay shows high correlation to the cell infectivity
assay (TCID50) with a minimum detection limit of 103 TCID50/ml (Saravanan et al., 2008).
d)
Nucleic acid recognition methods
Reverse transcription PCR (RT-PCR) techniques based on the amplification of parts of the N and F protein
genes has been developed for the specific diagnosis of PPR (Couacy-Hymann et al., 2002; Forsyth &
Barrett, 1995). This technique is 1000 times more sensitive than classical virus titration on Vero cells
(Couacy-Hymann et al., 2002) with the advantage that results are obtained in 5 hours, including the RNA
extraction, instead of 10–12 days for virus isolation. A multiplex RT-PCR, based on the amplification of
fragments of N and M protein genes, has been reported (George et al., 2006). Another format of the N genebased RT-PCR has also been described (Saravanan et al., 2004; Kumar et al., 2007). Instead of analysing
the amplified product – the amplicon – by agarose gel electrophoresis, it is detected on a plate by ELISA
through the use of a labelled probe. This RT-PCR-ELISA is ten times more sensitive than the classical RTPCR. In recent years, nucleic acid amplification methods for PPR diagnosis have been significantly
improved with quantitative real-time RT-PCR (Adombi et al., 2011; Balamurugan et al., 2010; Bao et al.,
2008; Batten et al., 2011; Kwiatek et al., 2010). This method is also ten times more sensitive than the
conventional RT-PCR, as well as minimising the risk of contamination. The application of nucleic acid
isothermal amplification to PPR diagnosis has also been described (Li et al., 2010). The sensitivity of this
assay seems to be similar to that of the real-time RT-PCR. This assay is simple to implement, rapid and the
result can be read by naked eye.
Because this is a rapidly developing field, users are advised to contact the OIE and FAO1 Reference
Laboratories for PPR (see Table given in Part 4 of this Terrestrial Manual) for advice on the most
appropriate techniques.
e)
Culture and isolation methods
Even when diagnosis has been carried out by rapid techniques, the virus should always be isolated from
field samples in tissue cultures for further studies (Durojaiye et al., 1983; Lefevre & Diallo, 1990).
PPRV may be isolated in primary lamb kidney/lung cells and some cell lines (Vero, B95a). Unfortunately,
PPRV isolation using such cells is not always successful on first passage and may require multiple blind
passages. Recently derivatives of cell lines (Vero, CV1) expressing the morbillivirus receptor, the signalling
lymphocyte activation molecule (SLAM or CD150), have been developed that can enable isolation of field
viruses from pathological specimens in less than 1 week, without requirement for blind passages. These
include a derivative of the monkey cell line CV1 expressing goat SLAM (Adombi et al., 2011) and derivatives
of Vero cells expressing dog SLAM. Monolayer cultures are inoculated with suspect material (swab material,
buffy coat or 10% tissue suspensions) and examined daily for evidence of cytopathic effect (CPE). The CPE
produced by PPRV can develop within 5 days and consists of cell rounding and aggregation culminating in
syncytia formation in lamb kidney cells and cell lines expressing SLAM. In Vero cells, it is sometimes difficult
to see the syncytia. If they exist, they are very small. However, small syncytia are always seen in infected
Vero cells stained with haematoxylin and eosin. Syncytia are recognised by a circular arrangement of nuclei
giving a ‘clock face’ appearance. Cover-slip cultures may give a CPE earlier than day 5. Some cells may
contain intracytoplasmic and intranuclear inclusions, others may be vacuolated. Similar cellular changes
may be seen in stained histopathological sections of infected tissues. After 5–6 days, blind passages should
always be carried out as CPE may take time to appear.
3.
Serological tests
The demonstration of antibodies in PPRV infected goats and sheep can be used to support a diagnosis by the
antigen-detection tests. Tests that are routinely used are the virus neutralisation (VN) test and the competitive
ELISA.
1
Food and Agriculture Organization of the United Nations.
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Chapter 2.7.11. — Peste des petits ruminants
a)
Virus neutralisation (the prescribed test for international trade)
This test is sensitive and specific, but it is time-consuming. The standard neutralisation test is now usually
carried out in 96-well microtitre plates (Rossiter et al., 1985) although roller-tube cultures may also be used.
Vero cells are preferred, but primary lamb kidney cells may also be used.
This test requires the following materials: cell suspensions at 600,000/ml; 96-well cell culture plates; sera to
be titrated (inactivated by heating to 56°C for 30 minutes); complete cell culture medium; PPRV diluted to
give 1000, 100, 10 and 1 TCID50/ml.
b)
i)
Dilute the sera at 1/5, and then make twofold serial dilutions in cell culture medium.
ii)
Mix 100 µl of virus at 1000 TCID50/ml (to give 100 TCID50 in each well) and 100 µl of a given dilution of
serum (using six wells per dilution) in the wells of the cell culture plate.
iii)
Arrange a series of control wells for virus and uninfected cells as follows: six wells with 100 TCID50
(100 µl) per well; six wells with 10 TCID50 (100 µl) per well; six wells with 1 TCID50 (100 µl) per well; six
wells with 0.1 TCID50 (100 µl) per well; and six wells with 200 µl of virus-free culture (control cells) per
well.
iv)
Make the wells containing the virus controls up to 100 µl with complete culture medium, and incubate
the plates for 1 hour at 37°C.
v)
Add 50 µl of cell suspension to each well. Incubate the plates at 37°C in the presence of CO2.
vi)
Read the plates after 1 and 2 weeks of incubation. The results should be as follows: 100% CPE in virus
control wells of 100 and 10 TCID50, 50% CPE for the 1 TCID50 dilution, no CPE for the 0.1 TCID50
dilution, no CPE in wells where the virus had been neutralised by serum during the test, and CPE in
wells where the virus had not been neutralised by serum during the test.
Competitive enzyme-linked immunosorbent assay
Several competitive ELISAs (C-ELISA) have been described, based on the use of MAbs that recognise virus
proteins. They are of two types: those where the MAb recognises the N protein and use recombinant N
protein produced in baculovirus as the antigen (Choi et al., 2005; Libeau et al., 1995); and those with a viral
attachment protein (H) specific MAb and antigen consisting of purified or part-purified PPRV (vaccine strain)
(Anderson & McKay, 1994; Saliki et al., 1993; Singh et al., 2004b). All the assays work on the principle that
antibodies to PPRV in test sera can block the binding of the MAb to the antigen.
Advice on the use and applicability of ELISA methods is available from the OIE Reference Laboratories for
PPR. Some methods are available as commercial kits. An example protocol for one method (Libeau et al.,
1995) is given below.
i)
Coat microtitre plates (such as Nunc Maxisorp) with 50 µl of a predetermined dilution of N-PPR protein
(produced by a recombinant baculovirus) for 1 hour at 37°C with constant agitation.
ii)
Wash the plates three times and blot dry.
iii)
Distribute 45 µl of blocking buffer (PBS + 0.05% Tween 20 + 0.5% fetal calf serum) to all wells, and
then add 5 µl of test sera to test wells in duplicate (at a final dilution of 1/20) and 5 µl of the different
control sera (strong positive, weak positive and negative serum) to control wells.
iv)
Add 50 µl of MAb diluted to working strength as advised by the supplier, and incubate at 37°C for
1 hour.
v)
Wash the plates three times and blot dry.
vi)
Add 50 µl of anti-mouse conjugate diluted 1/1000, and incubate at 37°C for 1 hour.
vii)
Wash the plates three times.
viii) Prepare OPD in hydrogen peroxide solution. Add 50 µl of substrate/conjugate mixture to each well.
Stop the reaction after 10 minutes with 50 µl of 1 M sulphuric acid.
ix)
Read on an ELISA reader at 492 nm.
The absorbance is converted to percentage inhibition (PI) using the formula:
PI = 1 – (absorbance of the test wells/absorbance of the MAb control wells) × 100
Sera showing PI greater than 50% in both duplicate wells are positive.
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Chapter 2.7.11. — Peste des petits ruminants
C. REQUIREMENTS FOR VACCINES
1.
Background
a)
Rationale and intended use of the product
Sheep and goats that recover from PPR develop an active life-long immunity against the disease (Durojaiye,
1982). Several homologous PPR vaccines are available, being cell culture-attenuated strains of natural
PPRV (Sen et al., 2010). In 1998, the OIE International Committee endorsed the use of such a vaccine in
countries that have decided to follow the ‘OIE pathway’ for epidemiological surveillance for rinderpest in
order to avoid confusion when serological surveys are performed. There have also been three published
reports on the preliminary results from recombinant capripox-based PPR vaccines that are able to protect
against both capripox and PPR (Berhe et al., 2003; Diallo et al., 2002, Chen et al., 2010). The production
and validation of vaccine from the commercially available attenuated PPRV is described here.
2.
Outline of production and minimum requirements for conventional vaccines
a)
Characteristics of the seed
The PPRV vaccine Nigeria 75/1 strain is a live vaccine cultured in Vero cells. The original strain of the virus
was isolated in Nigeria in 1975 (Taylor & Abegunde, 1979). It has been attenuated by serial passages in
Vero cell cultures (Diallo et al., 1989b). The strain provided for vaccine production is the 70th passage in
Vero cells (PPRV 75/1 I.K6 BK2 Vero 70). It is stored in freeze-dried form at –20°C and may be obtained
from Reference Laboratories (see Table given in Part 4 of this Terrestrial Manual). Tests of vaccine activity
show that it retained the ability to protect (at a dose of 103 TCID50) up to the 120th passage in Vero cells, the
latest passage tested so far.
b)
Method of culture
•
Cells
PPR vaccine is produced in Vero cells, which must be free from all bacterial, fungal and viral contamination.
•
Culture medium
The culture medium consists of minimal essential medium (MEM) supplemented with antibiotics (for
example penicillin + streptomycin at final concentrations of 100 IU [International Units]/ml and 100 µg/ml,
respectively), and an antifungal agent (nystatin [Mycostatin] at a final concentration of 50 µg/ml). The
medium is enriched with 10% fetal calf serum (complete medium) for cell growth. This proportion of serum is
reduced to 2% for maintenance medium when the cell monolayer is complete.
•
Primary seed batch of vaccine virus
This consists of virus in its 70th passage in Vero cells (PPRV 75/1 LK6 BK2 Vero 70). The freeze-dried
contents of a flask from the seed bank are reconstituted with 2 ml of sterile water (or cell culture medium
without serum). This liquid is mixed with Vero cells suspended in complete culture medium to provide at
least 0.001 TCID50 per cell. Cell culture dishes are filled with this virus/cell mixture (around 2 × 107 Vero
cells in a 175 cm2 dish), and are incubated at 37°C. The cultured cells are examined regularly to detect a
CPE. The medium is renewed every 2 days, reducing the proportion of serum to 2% once the cell monolayer
is complete. Virus is first harvested when there is 40–50% CPE. This viral suspension is stored at –70°C.
Successive harvesting is made every 2 days until the CPE reaches 70–80%, which is the time for final
freezing of the culture dishes (in general, at least two further harvestings can be made before final freezing
of the culture dishes). All suspensions of virus collected are submitted to two freeze–thaw cycles, then
added to form a single batch, which serves as the primary seed batch. This is divided into small volumes in
bottles and stored at –70°C. The contents of five flasks are thawed and titrated (minimum titre required:
105 TCID50/ml). It is best to freeze-dry this seed in order to store it at –20°C. In this case it will be necessary
to titrate the freeze-dried virus (five bottles). A batch made up in this way must pass all tests for sterility.
When preparing seed batches, it is important to avoid infecting the cells with high doses of virus (high
multiplicity of infection), as this will lead to accumulation of defective particles in the viral suspension
produced, which will diminish the titre of subsequent products. On the other hand, very weak multiplicity of
infection (e.g. 0.0001) will prolong the culture time.
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Chapter 2.7.11. — Peste des petits ruminants
•
Preparation of the working seed batch
This is done under the same conditions as for the primary seed batch. A large stock of virus is formed, from
which the final vaccine will be produced. This batch is distributed into receptacles and stored at –70°C. It
must satisfy tests for sterility. Five samples are titrated (minimum titre required: 106 TCID50/ml).
c)
Validation as a vaccine
It is necessary to confirm or rule out the presence of PPRV in the product under test. For this purpose, antiPPR serum is used to neutralise the virus in cell culture.
•
Test procedure
i)
Mix the contents of two vaccine bottles with sterile double-distilled water to provide a volume equal to
the volume before freeze-drying.
ii)
Make tenfold dilutions of the reconstituted vaccine in serum-free culture medium (0.5 ml of viral
suspension + 4.5 ml of medium).
iii)
Make two series of mixtures for virus dilutions from each bottle on a 96-well plate as follows:
Series 1:
Dilutions of viral suspension:
Viral suspension (in µl)
Culture medium (in µl)
–1
50
50
–2
50
50
–3
50
50
–4
50
50
Series 2:
Dilutions of viral suspension:
Viral suspension (in µl)
PPR antiserum (in µl)
–1
50
50
–2
50
50
–3
50
50
–4
50
50
(Note: PPR antiserum used for this purpose is prepared in goats and freeze-dried. It is reconstituted
with 1 ml of sterile double-distilled water in a dilution of 1/10.)
iv)
Incubate the mixtures at 37°C for 1 hour
v)
Add to each well 100 µl of cells suspended in complete culture medium (30,000 cells/well).
vi)
Incubate the microplate at 37°C in the presence of CO2.
vii)
Read the plate after 10–15 hours of incubation.
Normally a CPE is present only in the wells containing cells infected with the mixture of virus and culture
medium. If it is detected in the wells of Series 2, it will be necessary to identify PPRV by
immunofluorescence, using a PPR MAb, or by immunocapture (specific PPR MAb, and the immunocapture
test kit are available from the OIE Reference Laboratory for PPR in France [see Table given in Part 4 of this
Terrestrial Manual]). If this identification confirms the presence of PPRV, the PPR antiserum used must have
been too weak, or the batch must be changed. If immunofluorescence or immunocapture is negative, a viral
contaminant must be present, and the material under test must be destroyed.
2.
Method of manufacture
a)
Vaccine production
This operation is performed on a larger scale. Cells can be infected with virus at a multiplicity of infection as
before or with high doses, e.g. up to 0.01. Products of the various harvests, after two freeze–thaw cycles,
are brought together (to form the final product) and stored at –70°C pending the results of titration and tests
for sterility. If these results are satisfactory, the vaccine is freeze-dried.
b)
Freeze-drying
The freeze-drying medium (Weybridge medium) is composed of 2.5% (w/v) lactalbumin, 5% (w/v) sucrose
and 1% (w/v) sodium glutamate, pH 7.2.
This medium is added to an equal volume of viral suspension for freeze-drying (which may have been
diluted beforehand to provide the desired number of vaccine doses per bottle). The resulting mixture is kept
cool, homogenised, then distributed into bottles and freeze-dried. At the end of a freeze-drying cycle, the
probe is adjusted and kept at 35°C for 4 hours. Once this operation has been completed, the bottles are
capped under vacuum. Randomly selected samples (e.g. 5%) of this final batch are submitted to tests for
innocuousness, efficacy and sterility, and residual moisture is estimated by the Karl Fisher method (optimum
≤3.5%). If the tests give unsatisfactory results, the entire batch is destroyed.
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Chapter 2.7.11. — Peste des petits ruminants
3.
In-process control
Cells used in cultures must be checked for normal appearance and shown to be free from contaminating viruses,
especially bovine viral diarrhoea virus. A virus titration must be undertaken on the seed lot: using MEM (serumfree) medium, a series of tenfold dilutions is made (0.5 ml virus + 4.5 ml diluent) down to 10–6 of the product to be
titrated. Vero cells from one flask are trypsinised and suspended in complete culture medium at 300,000/ml. They
are distributed on a 96-well plate (30,000 cells per well, equivalent to 100 µl of cell suspension). Then, 100 µl of
virus diluted tenfold is added to the cells (dilutions ranging from 10–2 to 10–6). One row of wells serves as a
control for uninfected cells to which virus-free culture medium (100 µl) is added. The plate is incubated at 37°C in
the presence of CO2. The plates are read (by examining for CPE) 10–15 days after infection.
Virus titre is determined by the Spearman–Kärber method. The minimum titre per dose is 102.5.
4.
Batch control
a)
Identity
The contents of one container from each filling lot must be checked for identity by culture after neutralisation
with specific antiserum.
b)
Sterility
This consists of testing for viral, bacterial or fungal contaminants. It is done on cells and sera before their
use in vaccine production, and on the seed stock and the vaccine before and after freeze-drying. Any
product that fails this test for sterility is destroyed.
Tests for sterility and freedom from contamination of biological materials may be found in chapter 1.1.7.
c)
Safety
This test is done in rodents in order to detect any nonspecific toxicity associated with the product. The test
requires reconstituted vaccine in solvent (mixed contents of five bottles), six guinea-pigs, each weighing
200–250 g; ten unweaned mice (17–22 g, Swiss line or similar).
Vaccine, 0.5 ml, is injected intramuscularly into a hind limb of two guinea-pigs, 0.5 ml into the peritoneal
cavity of two guinea-pigs, and 0.1 ml into the peritoneal cavity of six mice. Two guinea-pigs and four mice
are kept as uninoculated controls. The animals are observed for 3 weeks. If one guinea-pig or two mice die,
the test must be repeated. Dead animals undergo post-mortem examination to ascertain the cause of death.
At the end of 3 weeks of observation, all animals are killed for post-mortem examination. All the results are
recorded. The vaccine is considered to be satisfactory if, during the first or second test, at least 80% of
animals remain in good health during the period of observation, and no significant post-mortem lesion is
found.
d)
Potency and efficacy in small ruminants
This test requires the following: vaccine reconstituted with normal saline (the mixed contents of five bottles)
to provide 100 doses and 0.1 dose/ml; six goats and six sheep, all approximately 1-year old and free from
antibodies to PPR; sterile syringes and needles; and pathogenic PPRV previously titrated in goats, diluted
with sterile normal saline to provide 103 of the 50% lethal dose for goats (LD50).
Vaccinate two goats and two sheep subcutaneously with 100 doses per animal; vaccinate two goats and two
sheep subcutaneously with 0.1 dose per animal; keep the remaining animals as in-contact controls. The
animals are observed and temperature measurements are recorded daily for 3 weeks. At the end of this
period, blood is taken from all animals for the preparation of sera. All animals are challenged by
subcutaneous injection of a 1 ml suspension of pathogenic PPRV (103 LD50 per animal). The animals are
observed and their body temperature measurements are recorded daily for 2 weeks.
The vaccine is considered to be satisfactory if all vaccinated animals resist the challenge infection, while at
least half of the in-contact controls develop signs of PPR. The serum neutralisation test must be positive for
PPR antibody (in serum diluted at least 1/10) in vaccinated animals only in samples taken 3 weeks after
vaccination. If any of the controls are also positive, the experiment must be repeated using another batch of
pathogenic PPRV. The batch of vaccine is destroyed if vaccinated animals react to the virulent challenge.
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Chapter 2.7.11. — Peste des petits ruminants
•
Titration of neutralising PPR antibody
This test requires the following: cell suspensions at 600,000/ml; 96-well cell culture plates; sera to be titrated
(inactivated by heating to 56°C for 30 minutes); complete cell culture medium; PPRV diluted to give 1000,
100, 10 and 1 TCID50/ml.
Dilute the sera at 1/5, then make a twofold serial dilutions in cell culture medium. Mix 100 µl of virus at
1000 TCID50/ml (to give 100 TCID50 in each well) and 100 µl of a given dilution of serum (using six wells per
dilution) in the wells of the cell culture plate. Arrange a series of control wells for virus and uninfected cells
as follows: six wells with 100 TCID50 (100 µl) per well; six wells with 10 TCID50 (100 µl) per well; six wells
with 1 TCID50 (100 µl) per well; six wells with 0.1 TCID50 (100 µl) per well; and six wells with 200 µl of virusfree culture (control cells) per well.
Make the wells containing the virus controls up to 100 µl with complete culture medium, and incubate the
plates for 1 hour at 37°C. Add 50 µl of cell suspension to each well. Incubate the plates at 37°C in the
presence of CO2. Read the plates after 1 and 2 weeks of incubation. The results should be as follows: 100%
CPE in virus control wells of 100 and 10 TCID50, 50% CPE for the 1 TCID50 dilution, no CPE for the
0.1 TCID50 dilution, no CPE in wells where the virus had been neutralised by serum during the test, and
CPE in wells where the virus had not been neutralised by serum during the test.
e)
Duration of immunity
Duration of immunity is at least 3 years.
f)
Stability
Freeze-dried vaccine can be kept for at least 2 years at 2–8°C (although storage at –20°C is better),
provided it is stored under vacuum and protected from light. Recently, it has been demonstrated that this
vaccine, suspended in medium containing trehalose and submitted to the ultra-rapid method of dehydration,
can resist at 45°C for a period of 14 days with minimal loss of potency (Worrwall et al., 2001).
5.
Tests on the final product
a)
Safety
See Section C.4.c.
b)
Potency
See Section C.4.d.
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ISMAIL T.M., HASSAN H.B., YOUSSEF N.M.A., AKHA G.M., ABD LE-HALIM M.M. & FATEHIA M.M. (1992). Studies on
prevalence of rinderpest and peste des petits ruminants antibodies in camel sera in Egypt. Vet. Med. J., 40, 49–
53.
KHALAFALLA A.I., SAEED I.K., ALI, Y.H., ABDURRAHMAN M B., KWIATEK O., LIBEAU G., OBEIDA A.A. & ABBAS Z. (2010).
An outbreak of peste des petits ruminants (PPR) in camels in the Sudan. Acta Trop, 116, 161–165.
KINNE J., KREUTZER R., KREUTZER M., WERNERY U. & WOHLSEIN P. (2010). Peste des petits ruminants in Arabian
wildlife. Epidemiol. Infect., 138, 1211–1214.
KWIATEK O., ALI Y.H., SAEED I.K., KHALAFALLA A.I., MOHAMED O.I., A.A. OBEIDA, ABDELRAHMAN M.B., OSMAN H.M.,
TAHA K.M., ABBAS Z., EL HARRAK M., LHOR Y., DIALLO A., LANCELOT R., ALBINA E. & LIBEAU G. (2011). Asian lineage
of peste des petits ruminants virus, Africa. Emerg. Infect. Dis., 17, 1223–1231.
KWIATEK O., KEITA D., GIL P., FERNANDEZ-PINERO J., JIMENEZ CLAVERO M.A., ALBINA E. & LIBEAU G. (2010).
Quantitative one-step real-time RT-PCR for the fast detection of the four genotypes of PPRV. J Virol. Methods,
165, 168–177.
KUMAR C.S., RAJ G.D., THANGAVELU A. & SHAILA M.S. (2007). Performance of RT-PCR-ELISA for the detection of
peste des petits ruminants virus. Small Rumin. Res., 72, 200–208.
LEFEVRE P.C. & DIALLO A. (1990). Peste des petits ruminants. Rev. sci. tech. Off. int. Epiz., 9, 951–965.
LEFEVRE P.C., DIALLO A., SCHENKEL F., HUSSEIN S. & STAAK G. (1991). Serological evidence of peste des petits
ruminants in Jordan. Vet. Rec., 128, 110.
LI L., BAO J., WU X., WANG Z., WANG J., GONG M., LIU C. & LI J. (2010). Rapid detection of peste des petits
ruminants virus by a reverse transcription loop-mediated isothermal amplification assay. J. Virol. Methods, 170
(1–2), 37–41.
LIBEAU G., DIALLO A., COLAS F. & GUERRE L. (1994). Rapid differential diagnosis of rinderpest and peste des petits
ruminants using an immunocapture ELISA. Vet. Rec., 134, 300–304.
LIBEAU G., PREHAUD C., LANCELOT R., COLAS F., GUERRE L., BISHOP D.H.L. & DIALLO A. (1995). Development of a
competitive ELISA for detecting antibodies to the peste des petits ruminants virus using a recombinant
nucleoprotein. Res. Vet. Sci., 58, 50–55.
MAJIYAGBE K.A., NAWATHE D.R. & ABEGUNDE A. (1984). Rapid diagnosis of PPR infection, application of immunoelectro-osmophoresis (IEOP) technique. Rev. Elev. Med. Vet. Pays Trop., 37, 11–15.
MORNET P., ORUE J., GILBERT Y., THIERY G. & SOW M. (1956). La peste des petits ruminants en Afrique Occidentale
Française. Ses rapports avec la peste bovine. Rev. Elev. Med. Vet. Pays Trop., 9, 313–342.
OZKUL A., AKCA Y., ALKAN F., BARRETT T., KARAOGLU T., DAGALP S.B., ANDERSON J., YESILBAG K., COKCALISKAN C.,
GENCAY A. & BURGU I. (2002). Prevalence, distribution, and host range of Peste des petits ruminants virus in
Turkey. Emerg. Infect. Dis., 8, 708–712.
PERL S., ALEXANDER A., YAKOBSON B., NYSKA A., HARMELIN A., SHEIKHAT N., SHIMSHONY A., DAVIDSON N., ABRAMSON
M. & RAPOPORT E. (1994). Peste des petits ruminants (PPR) of sheep in Israel: case report. Israel J. Vet. Med.,
49, 59–62.
ROGER F., GUEBRE YESUS M., LIBEAU G., DIALLO YIGEZU L.M. & YILMA, T. (2001). Detection of antibodies of
rinderpest and peste des petits ruminants viruses (Paramyxoviridae, Morbillivirus), during a new epizootic disease
in Ethiopian camels (Camelus dromedarius). Rev. Med. Vet., 152, 265–268.
ROGER F., YIGEZU L.M., HURARD C., LIBEAU G., MEBRATU G.Y., DIALLO A. & FAYE B. (2000). Investigations on a new
pathological condition of camels in Ethiopia. J. Camel Pract. Res., 7, 163–165.
ROSSITER P.B., JESSETT D.M. & TAYLOR W.P. (1985). Microneutralisation systems for use with different strains of
peste des petits ruminants virus and rinderpest virus. Trop. Anim. Health Prod., 17(2), 75–81.
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SALIKI J.T., LIBEAU G., HOUSE J.A., MEBUS C.A. & DUBOVI E.J. (1993). Monoclonal antibody-based blocking enzymelinked immunosorbent assay for specific detection and titration of peste-des-petits ruminants virus antibody in
caprine and ovine sera. J. Clin. Microbiol., 31, 1075–1082.
SARAVANAN P., SEN A., BALAMURUGAN V., BANDYOPADHYAY S.K. & SINGH R.K. (2008). Rapid quality control of a live
attenuated Peste des petits ruminants (PPR) vaccine by monoclonal antibody based sandwich ELISA.
Biologicals, 36, 1–6.
SARAVANAN P., SINGH R.P., BALAMURUGAN V., DHAR P., SREENIVASA B.P., MUTHUCHELVAN D., SEN A., ALEYAS A.G.,
SINGH R.K. & BANDYOPADHYAY S.K. (2004). Development of an N gene-based PCR-ELISA for detection of Pestedes-petits-ruminants virus in clinical samples. Acta Virol., 48, 249–255.
SEN A., SARAVANAN P., BALAMURUGAN V., RAJAK K. K., SUDHAKAR S. B., BHANUPRAKASH V., PARIDA S. & SINGH R. K.
(2010). Vaccines against peste des petits ruminants virus. Expert Rev. Vaccines, 9 (7), 785–796.
SHAILA M.S., PURUSHOTHAMAN V., BHASAVAR D., VENUGOPAL K. & VENKATESAN R.A. (1989). Peste des petits
ruminants in India. Vet. Rec., 125, 602.
SINGH R.P., SREENIVAS B.P., DHAR P. & BANDYOPADHYAY S.K. (2004a) .A sandwich-ELISA for the diagnosis of
Peste des petits ruminants (PPR) infection in small ruminants using anti-nucleocapsid protein monoclonal
antibody. Arch. Virol., 149, 2155–2170.
SINGH R.P., SREENIVASA B.P., DHAR P., SHAH L.C. & BANDYOPADHYAY S.K. (2004b). Development of a monoclonal
antibody based competitive-ELISA for detection and titration of antibodies to peste des petits ruminants (PPR)
virus. Vet. Microbiol., 14, 3–15.
SWAI E. S., KAPAGA A., KIVARIA F., TINUGA D., JOSHUA G. & SANKA P. (2009). Prevalence and distribution of peste
des petits ruminants virus antibodies in various districts of Tanzania. Vet. Res. Commun,, 33, 927–936.
TAYLOR W.P. & ABEGUNDE A. (1979). The isolation of peste des petits ruminants virus from Nigerian sheep and
goats. Res. Vet. Sci., 26, 94–96.
TAYLOR W.P., ALBUSAIDY S. & BARRETT T. (1990). The epidemiology of peste des petits ruminants in the Sultanate
of Oman. Vet. Microbiol., 22, 341–352.
WANG Z., BAO J., WU X., LIU Y., LI L., LIU C., SUO L., XIE Z., ZHAO W. ZHANG W., YANG N., LI J., WANG S. & WANG J.
(2009). Peste des petits ruminants virus in Tibet, China. Emerg. Infect Dis., 15, 299–301.
WORRWALL E.E., LITAMOI J.K., SECK B.M. & AYELET G. (2001). Xerovac: an ultra rapid method for the dehydration
and preservation of live attenuated rinderpest and peste des petits ruminants vaccines. Vaccine, 19, 834–839.
*
* *
NB: There are OIE Reference Laboratories for Peste des petits ruminants
(see Table in Part 4 of this Terrestrial Manual or consult the OIE Web site for the most up-to-date list:
http://www.oie.int/en/our-scientific-expertise/reference-laboratories/list-of-laboratories/ ).
Please contact the OIE Reference Laboratories for any further information on
diagnostic tests, reagents and vaccines for Peste des petits ruminants
OIE Terrestrial Manual 2012
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CHAPTER 2.7.12.
SALMONELLOSIS
(S. abortusovis)
See Chapter 2.9.9. Salmonellosis
*
* *
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NB: Ve rsion a dopted by the Worl d A ssembly of De legates of the OIE in May 2009
CHAPTER 2.7.13.
SCRAPIE
SUMMARY
Scrapie is a naturally occurring, infectious, neurodegenerative disease of sheep and goats
characterised by vacuolar or spongy changes in the central nervous system (CNS). It has been
recognised as a clinical disorder for more than 250 years, and is now classified as a transmissible
spongiform encephalopathy (TSE), or prion disease, defined by the accumulation of an abnormal
form (referred to as PrPSc) of a host membrane glycoprotein (prion protein or PrP), in the CNS. In
some animals, PrPSc accumulation is also detectable in lymphoreticular tissues. Polymorphisms of
the sheep PrP gene are associated with susceptibility to scrapie. PrP genotyping has been used as
a tool in the control of classical scrapie, but no genotype appears to be completely resistant to
infection, and the recently identified variant – atypical scrapie – has been reported in sheep of PrP
genotypes that are apparently resistant to classical scrapie. Classical scrapie is endemic in many
parts of the world, where it has often been introduced by importation. Australia and New Zealand
have maintained freedom by use of strict restrictions on imports and other measures. The infection
in sheep may be passed from ewe to lamb in the period from parturition to weaning. Infection can
also pass horizontally to unrelated sheep or goats, especially when parturition occurs in confined
areas. Fetal membranes are known to be a source of infection, and sheep milk from clinically
affected animals has been demonstrated to transmit disease. The incubation time between primary
infection and clinical disease is nearly always longer than 1 year and may sometimes exceed the
commercial lifespan of the sheep. The majority of cases occur in sheep between 2 and 5 years of
age. Clinical disease only develops if the infection enters the CNS. Atypical scrapie, when it
presents clinically, appears to affect older animals predominantly, and occurs with a geographical
distribution suggestive of a spontaneous disease, although it can be transmitted experimentally.
Identification of the agent: The disease is recognised by the clinical signs, which are variable, but
generally start insidiously with behavioural abnormalities then progress to more obvious
neurological signs, including pruritus, incoordination and poor bodily condition. Clinical diagnosis is
confirmed by the presence of gray matter vacuolation of target areas within the brain tissue and/or
the immuno-detection of disease-specific accumulations of PrPSc in the brain or lymphoreticular
tissues. Immunochemical detection of the protein in brain samples forms the basis of rapid tests
used in active surveillance programmes. In experimental studies of the disease in sheep and goats,
detectable PrPSc accumulation in the brain does not start until several months after challenge, so it
must be remembered that a negative result does not necessarily equate to an unexposed animal.
Detection of PrPSc in lymphoreticular tissues during the incubation period of scrapie in some
animals offers a means of preclinical diagnosis of scrapie infection and may be particularly useful
for surveillance purposes. It can also be performed in vivo using biopsied tissue. It is not, however,
appropriate for atypical cases, or a proportion of classical cases, so can only be used to confirm
presence of infection and cannot be used to prove absence of disease.
Most, but not all, currently recognised forms of scrapie can be transmitted to laboratory rodents by
injecting them with infected brain tissue, but the variable efficiency of transmission coupled with
long incubation times preclude this as a practical diagnostic procedure.
Serological tests: Scrapie infection is not known to elicit any specific immune response and there
is no basis for establishing a diagnosis by detecting specific antibodies.
Requirements for vaccines and diagnostic biologicals: There are no biological products
available.
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Chapter 2.7.13. — Scrapie
A. INTRODUCTION
Scrapie (also known as la tremblante; Traberkrankheit; Gnubberkrankheit) is a naturally occurring progressive,
fatal, infectious, neurodegenerative disease of sheep and goats that has been recognised for at least 250 years,
and has been reported in Europe, North America, Asia and Africa. Scrapie and other related diseases in humans
and animals were later classified as transmissible spongiform encephalopathies (TSE), although they are now
known as prion diseases. The occurrence of scrapie preceded recognition of other prion diseases of mammals
and so, in retrospect, it is the archetype of prion disorders (Hörnlimann, 2006). Prion diseases are defined by the
accumulation of an abnormal isoform (PrPSc) of the host-encoded protein (PrPC) in the central nervous system
(CNS) in all cases. The abnormal protein may also be detected in other tissues such as in the lymphoreticular
system (LRS) and variably in other tissues/body fluids.
An atypical form of scrapie was first described in Norway in 1998 (Benestad et al., 2008). Subsequently, active
surveillance using rapid immunochemical methods has provided evidence for the widespread occurrence of this
atypical form of scrapie (also known as Nor98), throughout Europe, with reports of cases also in the Falkland
Islands (Epstein et al., 2005) and North America (Loiacono et al., 2009). Although the epidemiology is not
suggestive of transmission in the field (Benestad et al., 2008) this type of scrapie can be transmitted
experimentally (Simmons et al., 2007) Retrospective studies have now identified British cases from the late
1980s, predating active surveillance (EFSA, 2005b). Atypical scrapie has been identified in sheep of genotypes
considered to be resistant to classical scrapie, and in goats (Arsac et al., 2007; Seuberlich et al., 2007).
The interaction of agent variables (particularly strain) and host variables determines the disease phenotype. In
sheep, different PrP genotypes are associated with relative susceptibility to TSEs (Goldmann, 2008).
Polymorphisms at codon 171 are of particular significance in determining overall susceptibility of sheep to
classical scrapie, while variations at 141 and 154 affect susceptibility of sheep to the more recently identified
atypical form of disease (Benestad et al., 2008; Moum et al., 2005; Saunders et al., 2006).
Characterisation of different strains of scrapie isolates has, historically, relied upon transmission to rodents
(biological strain typing) (Bruce et al., 2004), principally using inbred (wild-type) mice, but increasingly also a
number of different transgenic mouse constructs (Groschup & Buschmann, 2008). Molecular TSE-typing uses
differential epitope binding of PrPSc in immuno-histochemistry (IHC) or Western immunoblotting (Jeffrey et al.,
2001). The ability to distinguish scrapie from bovine spongiform encephalopathy (BSE) is of particular importance
in small ruminants because of the zoonotic nature of the latter and the potential for past exposure of small
ruminants to the feed-borne BSE agent. However, the mechanisms by which strain and host parameters influence
disease phenotype are still incompletely understood.
Because of the known inadequacies of baseline (passive) surveillance and the absence of active surveillance
components, the true scrapie status of many countries is unknown, and it is probably impossible to establish
freedom from infection in a national flock without recourse to disproportionate levels of active surveillance. Some
countries have never recorded the disease against a background of general and/or targeted surveillance, while
others have maintained freedom for various periods through rigorous preventative policies and monitoring. The
disease usually occurs in sheep 2–5 years of age. Rarely are cases present in sheep less than 1 year of age. In
atypical scrapie, significant numbers of cases have been reported in sheep over 5 years of age. In some
instances, the commercial lifespan of the sheep may be too short or exposure has occurred too late in life to allow
the clinical disease to develop. Classical scrapie has also been described in goats, and captive moufflon (Ovis
musimon), a primitive type of sheep. Most breeds of sheep are affected. The infection in sheep may be passed
from ewe to lamb in the period from parturition to weaning. Infection can also pass horizontally to unrelated sheep
or goats, especially when parturition occurs in confined areas. Fetal membranes are known to be a source of
infection, and milk has recently been shown to be infectious to lambs (Konold et al., 2008). Pasture previously
grazed by, or buildings previously inhabited by, infected sheep may also represent a risk. Animals incubating the
disease, and even animals that never develop clinical signs, may still be a source of infection to others.
The biohazard for humans from scrapie diagnostic testing appears to be limited, but appropriate precautions
should be taken. Creutzfeldt-jakob disease (CJD) has been found to occur at no greater frequency in those with
occupations providing closest contact with the agent, but the extreme chemical and physical resistance of the
scrapie agent and the fact that it is experimentally transmissible by injection to a wide spectrum of mammalian
species suggest the prudence of preventing human exposure.
B. DIAGNOSTIC TECHNIQUES
1.
Identification of the agent
The specific causative agent has never been isolated or described for any prion disease. Characterisation is
therefore based on the identification of phenotypic parameters, such as clinical signs, through histopathological
profiles, to increasingly complex immunochemical and biological parameters.
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Chapter 2.7.13. — Scrapie
The clinical signs of scrapie (Ulvund, 2006) usually start insidiously, often with behavioural changes that are
evident only from repeated inspections. These subtle presenting features, which may include apparent confusion,
separation from the flock and a staring gaze, progress to a more definite neurological illness, frequently
characterised by signs of pruritus and ataxia or incoordination of gait. Either the pruritus or the ataxia usually
emerges to dominate the clinical course. Death may occur after a protracted period of only vague neurological
signs or may even occur without premonitory signs. These clinical signs, individually, are not disease-specific and
clinical suspicion of disease should be confirmed by further testing.
Pruritus is recognised principally by compulsive rubbing or scraping against fixed objects, nibbling at the skin and
scratching with a hind foot or horns. This results in extensive loss of wool, particularly over the lateral thorax,
flanks, hindquarters and tail head. The persistence of pruritus often results in localised self-inflicted skin lesions.
These may occur in areas of wool loss and on the poll, face, ears and limbs. A characteristic ‘nibble reflex’ can
often be elicited by scratching the back, and may also be evoked by the sheep’s own scraping movements. Some
sheep or goats with scrapie, however, may not present with evident signs of pruritus.
Ataxia or incoordination of gait may first become apparent as difficulty in positioning the hind limbs on turning,
swaying of the hindquarters and a high stepping or trotting gait of the forelimbs. Stumbling and falling occur, but
the sheep is generally able to quickly regain a standing posture. These signs progress to weakness and
recumbency. Information on the phenotypic variants of scrapie, termed atypical scrapie, or Nor98 suggests
clinical features being dominated by ataxia in the absence of pruritus; circling has also been observed (Benestad
et al., 2003; Konold et al., 2007; Moum et al., 2005; Simmons et al., 2009; Ulvund, 2006). Other signs of scrapie
may include teeth grinding (bruxism), low head carriage, a fine head or body tremor and, rarely, seizures or visual
impairment. In most cases, there is also a loss of bodily condition or weight.
Progress of the clinical disease is very variable, lasting for a week or up to several months, with an inevitably fatal
outcome. There is also variation in the clinical signs among individual animals and in different breeds of sheep.
These variations may be due to the influence of host genotype and strain of agent. Environmental factors may
also have an influence on the disease course. The clinical diagnosis of individual cases of scrapie can therefore
be difficult. The clinical signs may, especially in the early phase of the disease, resemble those of some other
conditions of adult sheep, including ectoparasitism, pseudorabies (Aujeszky’s disease), rabies, cerebral listeriosis,
ovine progressive pneumonia (maedi-visna), pregnancy toxaemia (ketosis), hypomagnesaemia and chemical and
plant intoxications.
Video-clips illustrating the clinical signs of scrapie can be viewed on the website of the OIE Reference Laboratory
at Animal Health and Veterinary Laboratories Agency (AHVLA), which is also the European Commission TSE
Community Reference Laboratory1. Other historical videotape footage of classical scrapie signs can also be
sourced (Wells & Hawkins, 2004). Video-clips of atypical scrapie signs are also available (Konold et al., 2007).
Full narratives describing the clinical signs of scrapie are available in the literature (Benestad et al., 2008; Konold
et al., 2007; Ulvund, 2006).
According to the prion hypothesis, demonstration of PrPSc would constitute identification of the agent, but, by
definition, transmission from infected tissues, usually to laboratory rodents by injection, is the only available
means of detection of infectivity. Long incubation periods and the failure of some natural scrapie sources to
transmit to specific mice strains mean that it is impractical to use the criterion of transmissibility for diagnosis.
However, biological characterisation on transmission is an important experimental component of the definition of
any emerging new phenotypic variants of scrapie and for discriminatory approaches to distinguish cases of
scrapie from BSE in sheep or goats (see footnote 1). Given the absence of any specific gross pathological
changes, the laboratory diagnosis of classical scrapie (Gavier-Widen et al., 2005) has, in the past, been reliant on
the demonstration of histopathological changes, notably vacuolation, in the CNS. Prior to the routine use of
immunochemical detection of PrPSc, morphological demonstration of PrPSc in the form of scrapie-associated
fibrils (SAF) was employed as an adjunct to histopathological diagnosis. SAF are visualised in unfixed brain
extracts using negative-stain electron microscopy and have also been recovered from formaldehyde-fixed and
autolysed brain tissue (Chaplin et al., 1998; Stack et al., 1996).
The histopathological diagnosis was historically based on examination of a single section of medulla oblongata
taken at the level of the obex, considered the predilection site for morphological vacuolar changes (Wood et al.,
1997). This approach is still valid for the confirmation of classical scrapie, but it will not detect atypical scrapie.
However, PrPSc detection using IHC examination and/or Western immunoblotting techniques, performed on the
medulla oblongata, have increased the efficiency of the detection methods, and this has now extended to the
active surveillance of large populations using rapid PrP detection tests (see below). Detectable PrPSc precedes
vacuolation and clinical signs, making the immuno-based tests a more sensitive option (Hamir et al., 2001). While
clinically suspect cases of scrapie should, where suitable samples are available, continue to be investigated
1
European Commission: TSE Community Reference Laboratory – Web Resources:
http://www.vla.gov.uk/science/sci_tse_rl_video.htm
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Chapter 2.7.13. — Scrapie
initially by histopathological examination for morphological changes, diagnostic criteria must now include the
demonstration of PrPSc in the CNS. The medulla remains the most consistent and appropriate diagnostic level of
the CNS for classical scrapie, however the accumulation pattern of PrPSc in atypical scrapie is different (Benestad
et al., 2008; Moore et al., 2008). In atypical scrapie, the medulla shows only minimal change, while much more
consistent and overt lesions can usually be identified in the cerebellum, thalamus and basal ganglia. Therefore,
taking practical and logistical sampling considerations into account the medulla and cerebellum should both be
examined as a minimum for robust diagnosis. This approach assists in the characterisation of the prion protein
that is present, and contributes to discrimination of disease phenotypes, particularly between classical and
atypical scrapie, and indeed is also sufficient for the discrimination of scrapie and BSE (Baron et al., 2006).
Some commercially available rapid methods for the detection of PrPSc, introduced originally for the diagnosis of
BSE, are also approved for scrapie diagnosis (European Food Safety Authority [EFSA] (2005a). These rapid tests
take the form of Western immunoblot, luminescence (LIA)- or enzyme-linked immunosorbent assay (ELISA)based methods, and provide preliminary screening from which positive or inconclusive results are subject to
examination by confirmatory IHC or Western blot methods. All these methods have been shown to be able to
detect classical scrapie in the appropriate samples (EFSA, 2005a), but some have subsequently been shown to
detect atypical scrapie to a varying degree or not at all. The analytical sensitivity of these kits is kept under review
by the European Commission, and links to information on the performance of currently approved tests can be
found on the Reference Laboratory webpages (see footnote 1).
Failure to observe characteristic histological changes or to detect disease-specific PrP/SAF does not confirm the
absence of the disease; agreement between the results of multiple diagnostic approaches provides the best
assurance of accuracy. Clearly, in surveillance situations where monitoring is for the purpose of obtaining
evidence of freedom from scrapie in small ruminant species, it may be necessary to apply multiple diagnostic
criteria and to use at least two laboratory methods (histopathological and IHC, or immunoblotting) on accurately
sampled CNS tissue (minimum medulla and cerebellum) to maintain a high degree of confidence in negative
results.
Passive surveillance of scrapie, comprising the examination of CNS material from clinically suspect cases, has, in
recent years, been complemented in many countries by active surveillance, targeting healthy adult culls and fallen
stock (diseased or dead animals, also called risk animals) screened at post-mortem using the rapid test methods.
In scrapie, the opportunity also exists for screening approaches that do not rely solely on examination of the CNS
tissue from dead animals to detect exposed animals, but uses the widespread presence of PrPSc in
lymphoreticular tissue in many animals to enable demonstration of infected animals by biopsy of palatine tonsil,
nictitating membrane, superficial lymph nodes or, most recently, rectal mucosa lymphoreticular tissue (Gonzalez
et al., 2006). Any of these tissues can usefully be included in the screening of cull populations too, to maximise
disease detection. It must be noted that not all animals with classical scrapie have detectable lymphoreticular
involvement (Andreoletti et al., 2000; Gonzalez et al., 2006), and no PrPSc has yet been detected in the
lymphoreticular tissues of cases of sheep or goats with atypical scrapie (Benestad et al., 2003). However, the
testing of lymphoreticular tissue offers the opportunity to detect some animals infected with classical scrapie at
relatively early stages of incubation, before the CNS is positive.
Whereas large-scale testing, to determine freedom from scrapie, should include targeted examination of
peripheral tissues as well as the CNS of younger animals, surveillance for prevalence of disease could potentially
limit tissue examination to the CNS of adult sheep and goats for the reasons given above. However, testing to
estimate disease prevalence needs to take into account a number of factors, including the stratification of the
sheep-farming industry, dose or level of infection within particular flocks, frequency of disease and relative
involvement of the LRS in different genotypes, and the effect of genotype/agent strain combination on incubation
period.
The need to distinguish between cases of scrapie and possible BSE in sheep and goats has required the
development of diagnostic methods with the potential to discriminate between the agents causing these
infections. Studies suggest that the conformation of disease-specific PrP produced in BSE-infected sheep is
different from that of disease-specific PrP found in natural sheep scrapie (EFSA, 2005b; Jeffrey et al., 2001; Stack
et al., 2006). These conformational differences may be detected by immunoblotting or IHC techniques using
epitope-specific antibodies. Within the European Union, the strategy for this distinction comprises examination of
source CNS material after initial detection through active or passive surveillance (initial screening), in a primary,
secondary and tertiary phase procedure (see footnote 1), involving a Western immunoblot method capable of
such discrimination, followed by peer review and further investigation by biochemical and IHC methods, and
finally, if necessary, mouse transmission to a standard panel of wild-type mice (see Chapter 2.4.6 Bovine
spongiform encephalopathy). Interpretation of the in-vitro methods (Western immunoblot or ELISA) is reliant on
differences between BSE and scrapie in the N-terminal cleavage site for Proteinase-K digestion of PrPSc. The insitu IHC approach relies on distribution and epitope-specific labelling patterns of PrPSc in brain and
lymphoreticular tissues. Increasingly, for biological characterisation of agent strain, appropriate transgenic mouse
constructs are being used, although comprehensive data comparing models are still sparse.
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Chapter 2.7.13. — Scrapie
Quality control (QC), quality assurance (QA) and appropriate positive and negative control samples are essential
parts of testing procedures and advice and control materials can be requested from the OIE Reference
Laboratories.
a)
Sample preparation
Concerns regarding BSE in small ruminant populations and, the recognition of atypical scrapie, have
influenced the strategies for sampling and diagnosis. Although comprehensive sampling and multiple testing
methods would provide the most robust contingencies for these and possible future uncertainties in the
diagnosis of prion diseases of small ruminants, operational factors also determine what is practically and
economically possible. The relative implementation of passive and active surveillance programmes, and the
diagnostic methods applied, further influence sampling strategy. Selection and recommendation of methods
is therefore under constant review.
For routine diagnosis, the sampled CNS material is either stored fresh or frozen for subsequent biochemical
tests or is fixed for histological preparations. Where programmes are in place to identify possible infections
with BSE in small ruminant populations, all sampling should be conducted aseptically, using new sterile
disposable instruments, or instruments sterilised under conditions specified for the decontamination of prions
(see chapter 2.4.6). Cross contamination at necropsy/sampling should be avoided. Thus, in the following
procedures where fresh tissue is sampled for biochemical methods, an aliquot should, if required, be
reserved for transmission studies. Although in many instances disease can be confirmed on autolysed or
suboptimally preserved material, such samples can only provide limited evidence of the absence of scrapie.
Sheep in which clinical disease is suspected (passive surveillance) should be killed by intravenous
injection of barbiturate and the whole brain removed by standard necropsy procedures as soon after death
as possible. Whole brain removal is advisable to allow pathological examinations for differentiation between
possible different TSE phenotypes and differential diagnosis of non-TSE brain disorders. Methods of
subdividing the brain tissue, for application of PrP-detection techniques requiring fresh tissue and for
histological techniques, are dependent on the optimum sensitivities of each of the tests for different brain
areas and the compromise that precisely the same area cannot be used for both fresh/frozen and fixed
tissue approaches. The following protocol is suggested but may be subject to modification to satisfy the
particular portfolio of tests. Further information can be obtained from OIE Reference Laboratories (see
footnote 1 or Table in